Vol. 55, No. 4
MICROBIOLOGICAL REVIEWS, Dec. 1991, p. 586-620 0146-0749/91/040586-35$02.00/0 Copyright C 1991, American Society for Microbiology
Viruses and Viruslike Particles of Eukaryotic Algae JAMES L. VAN ETTEN,1* LESLIE C. LANE,' AND RUSSEL H. MEINTS2 Department of Plant Pathology, University of Nebraska, Lincoln, Nebraska 68583-0722,1 and Department of Botany and Plant Pathology, Center for Gene Research and Biotechnology, Oregon State University, Corvallis, Oregon 97331-2906 INTRODUCTION ...................................................................................
586
CHARACTERISTICS OF CHLORELLA SPECIES
587
......................................................................
Properties of Chlorella Species ...................................................................................587 590 Symbiotic Chlorella Species ................................................................................... Chlorella Strains NC64A, Nla, Pbi, and SAG-241-80 .................................................................591 General
DISCOVERY OF CHLORELLA VIRUSES ................................................................................. 591 Initial Observations
Development of
a
591
...................................................................................
Culturable
System
for
PBCV-1
...................................................................... 592
PROPERTIES OF CHLORELLA VIRUSES ................................................................................ 592 General Characteristics of
Composition
PBCV-1
of
PBCV-1
PBCV-1
594
Genome ...................................................................................
PBCV-1 Chlorella Strain NC64A Viruses Viruses Infecting Chlorella Strain Pbi REPLICATION OF PBCV-1 Attachment of PBCV-1 to Chlorella Strain NC64A Intracellular Site of PBCV-1 Replication Effect of PBCV-1 Infection Host DNA, RNA, and TRANSCRIPTION OF PBCV-1 Serology
592
...................................................................................
...................................................................................
of
596
597
...................................................................................
Additional
..............................................................................
598 598
...................................................................................
598
................................................................................... ....................................................................
598
.................................................................................
600
on
Protein
Syntheses .........................................604 604
...................................................................................
VIRUSES ENCODE DNA METHYLTRANSFERASES AND DNA SITE-SPECIFIC (RESTRICTION) ENDONUCLEASES
PBCV-1
607
...................................................................................
Methyltransferases and DNA Site-Specific Endonucleases ........................................607 Viral DNA Methyltransferases and DNA Site-Specffic Endonucleases ..............................607
DNA
Additional
SEQUENCE OF VIRUS-ENCODED DNA METHYLTRANSFERASES ............................................609
FUNCTION OF VIRUS-ENCODED DNA SITE-SPECIFIC ENDONUCLEASES AND DNA METHYLTRANSFERASES ...................................................................................
609
NATURAL HISTORY OF THE CHLORELLA VIRUSES ..............................................................610 COMPARISON OF CHLORELLA VIRUSES WITH OTHER VIRUSES ...........................................611
PROPERTIES OF VIRUSES OR VLPs OF OTHER ALGAE
615
ACKNOWLEDGMENTS ...................................................................................
616
REFERENCES ...................................................................................
616
photosynthesis, cell cycle regulation, cell motility, and nonMendelian (chloroplast) inheritance. Given the importance of eukaryotic algae, there is surprisingly little information about viruses or viruslike particles (VLP) in these organ-
INTRODUCTION
Eukaryotic algae range in size and complexity from small (microalgae) to large multicellular organisms (macroalgae), such as seaweeds. Most eukaryotic algae are free living, and members can grow and survive in many terrestrial and aquatic environments. A few, however, are found only in symbiotic association with other organisms such as molluscs, flatworms, coelenterates, protozoa, and fungi (lichens). Algae significantly influence aquatic environments, both as primary producers in the food chain and, conversely, as pollutants when growth becomes uncontrolled. Because most algae require only light, simple salts, and a nitrogen source for rapid growth, they are potential producers of proteins, carbohydrates, and fats for feeding humans and animals. In addition, algae are a source of useful compounds such as 13-carotene, alginic acid, carrageenans, and agar (12, 137, 148). Algae such as Chlamydomonas and Chlorella spp. have also served as models for studying unicellular organisms
*
612
........................................................
CONCLUDING COMMENTS ...................................................................................
isms. It is difficult to credit the first person who described a virus or VLP in a eukaryotic alga. The problem is complicated because early reports consisted solely of microscopic observations and, in some cases, cultures were not axenic. A few papers in the Russian literature as long as 30 years ago described a lytic activity in cultures of the green alga Chlorella pyrenoidosa (174, 224-226). This lytic activity was given the name "chlorellophage." However, these investigators acknowledged that their cultures were contaminated with bacteria. Furthermore, the chlorellophage had typical bacteriophage morphology (174, 224). Therefore, despite the name chlorellophage, the host for these particles is unclear. Several investigators, almost simultaneously, reported the presence of VLPs in eukaryotic algae in the early 1970s. Lee (73) described 50- to 60-nm polyhedral particles in vegetative cells of the red alga Sirodotia tenuissima, Pickett-Heaps
Corresponding author. 586
VOL. 55, 1991
(123) described polyhedral particles (240 nm in diameter) in germlings of the green alga Oedogonium sp., and Toth and Wilce (176) described 170-nm polyhedral particles in spores of the marine brown alga Chorda tomentosa. Since these initial reports, VLPs or viruses have been reported in at least 44 taxa of eukaryotic algae (Table 1). These are distributed in 10 of the 14 recognized classes of eukaryotic algae (141). Particles have not been found in any member of the families Euglenophyceae, Bacillariophyceae, Raphidophyceae, and Tribophyceae. VLPs and viruses of eukaryotic algae have been reviewed previously by Lemke (74), Sherman and Brown (149), Dodds (27, 28), Martin and Benson (84), and Van Etten et al. (190, 192). The review by Sherman and Brown contains several electron micrographs of these particles, including some not published elsewhere. One other report is noteworthy. In 1970 Schnepf et al. (143) described large (200 nm in diameter) polyhedral particles in an Aphelidium sp. (which, although described as a fungus, may actually be a protozoan) that parasitized the green alga Scenedesmus armatus. These particles were much larger than most fungal viruses (68) but were similar in size and morphology to VLPs subsequently found in eukaryotic algae. With few exceptions, the reports of VLPs in eukaryotic algae typically consist of single accounts of microscopic observations. Furthermore, many of them were with fieldcollected algae, which are not available for further study. Characterization of these particles was not pursued because of low concentrations. Several factors contributed to the low concentrations: (i) usually only a few algal cells contained particles, (ii) usually the cells contained particles only at one stage of the algal life cycle, (iii) usually cells containing particles did not lyse, and (iv) in most cases the particles were not infectious. In addition, some of the algae could not be cultured and others were inconvenient to work with (e.g., filamentous algae). However, these difficulties are no longer a problem for a family of viruses which infect and replicate in certain strains of unicellular Chlorella-like green algae. The first of the Chlorella viruses was discovered about 10 years ago in Chlorella-like algae (zoochlorella) symbiotic with Hydra viridis (96, 186, 187) and subsequently in Chlorella-like algae in Paramecium bursaria (132, 181, 183, 187). The algae from P. bursaria can be grown free of the paramecium, and the cultured Chlorella strains NC64A and Pbi or their equivalents serve as hosts for many viruses. These viruses can be produced in large quantities, assayed by plaque formation, and analyzed by standard bacteriophage techniques. This review will describe the biology of the Chlorella virus hosts, which normally exist as symbionts, and then focus on the discovery, biology, and molecular biology of the Chlorella viruses. Partially characterized viruses or VLPs of other eukaryotic algae are discussed at the end of the review. CHARACTERISTICS OF CHLORELLA SPECIES General Properties of Chlorella Species The genus Chlorella contains small spherical or ellipsoidal, unicellular, nonmotile, asexually reproducing green algae. Chlorella species have a rigid cell wall and typically contain a single chloroplast, which may or may not contain a pyrenoid. They have a simple developmental cycle and reproduce by mitotic division. Vegetative cells increase in size and divide into two, four, eight, or more progeny (called autospores), which are released by rupture or enzymatic
VIRUSES OF EUKARYOTIC ALGAE
587
digestion of the parental walls. The autospores develop rapidly into vegetative cells. The number of progeny formed from a single vegetative cell varies with species and environmental conditions (200). Chlorella species typically grow on a simple salts solution and utilize nitrate, ammonium, or amino nitrogen as a nitrogen source (116). However, they often grow best on casein hydrolysate (91, 151). Simple sugars such as sucrose or glucose increase the growth rate of some species but inhibit the growth of others. The doubling time varies with the strain or species, but many of them double in about 24 h. Algae assigned to the genus Chlorella are more heterogeneous than their simple morphology suggests (25, 58, 65, 151). Huss et al. are examining the relationships among Chlorella species by using DNA base ratios, DNA hybridization, and DNA reassociation kinetics (see reference 58 and references cited therein). The guanosine-plus-cytosine content of nuclear DNA, which ranges from 43 to 79% (49), reflects the heterogeneity of algae assigned to the genus Chlorella. However, most isolates assigned to the same species have similar G+C contents. The Chlorella base compositions were determined either by measuring DNA melting temperatures or by performing CsCl density equilibrium centrifugation. Recently, high levels of 5-methylcytosine (5mC) were found in the nuclear DNAs of Chlorella strain NC64A (21% of the cytosines were methylated [189]) and Chlorella sorokiniana (16% of the cytosines were methylated [22]). If high concentrations of 5mC are a common feature of Chlorella DNAs, many of the published Chlorella G+C contents may be low because 5mC can lower DNA melting temperatures and buoyant densities. The DNA content of Chlorella cells was reported to range from about 15 fg per cell for cultured Chlorella strains 211/le and NC64A to 47 fg per cell for symbiotic Chlorella strains isolated from hydras (88). We estimate that Chlorella strain NC64A contains about 40 fg (40 Mbp) of DNA per cell (180), which is almost three times higher than that found by McAuley and Muscatine (88). Assuming the higher value and assuming haploidy, the Chlorella strain NC64A genome is about twice the size of the Saccharomyces cerevisiae genome and one-half the size of the Arabidopsis thaliana genome. Repetitive sequences make up only a small fraction (about 5 to 8%) of the total Chlorella DNA (58). The cell wall composition of Chlorella species also varies widely (see, e.g., references 3, 76, 114, 171, 172, and 219). Typically, Chlorella cell walls contain polysaccharides with a variety of component sugars as well as lesser amounts of protein and lipid. In addition, walls of some Chlorella species contain a layer of sporopollenin (3), a polymer of carotenoid derivatives resistant to most chemicals (13). The difficulty of classifying Chlorella species is illustrated by a report that the chemical compositions of the cell walls of some isolates assigned to the same species differ markedly, e.g., for Chlorella vulgaris and Chlorella ellipsoidea (219). Because of the heterogeneity in the genus Chlorella, properties discovered in one Chlorella species or strain may not always apply to another. As noted below, viruses can distinguish at least two exsymbiotic Chlorella strains from each other and from other free-living Chlorella species. If viruses infecting additional Chlorella species are found, viruses could be useful in classifying Chlorella species in the same way that bacteriophages are useful in bacterial classification (see, e.g., reference 2).
588
VAN ETTEN ET AL.
MICROBIOL. REV. TABLE 1. Viruses and VLPs of eukaryotic algae
Host species'
Location of virogenic stroma
Infection and replication facts
Morphological
Nucleic acid
characteristics
Key references
Rhodophyceae
Porphyridium purpureumb
Unkc
Polyhedral, 40 nm in diameter
Unk
18
Unk
Polyhedral, 50-60 nm in diameter
Unk
73
Nucleus and cytoplasm
Unk
Two types of particles were observed in the same cell: (i) polyhedral, 99 nm in diameter; (ii) stalked particle, head 120 nm diameter, length 240 nm; surrounded by a membrane
Unk
124
Dinophyceae Gymnodinium uberrimunf *Gyrodinium resplendensg
Cytoplasm Cytoplasm
Unk Unk
Polyhedral, 385 nm in diameter Two types of particles: (i) tubular arrays, with particles 35 nm in diameter; (ii) particles 20 nm in diameter
Unk Unk
152 34, 162
Chrysophyceae Chromophysomonas cor-
Cytoplasm
Unk
Unk
127
Dinobryon sp.J
Cytoplasm Cytoplasm
Unk Unk
Two polyhedral particles, 50-60 nm and 150-180 nm, in diameter Polyhedral, 35 nm in diameter Polyhedral, 100 nm in diameter
Unk Unk
Hydrurus foetidusk Mallomonas sp. Paraphysomonas caelifri-
Cytoplasm Cytoplasm Cytoplasm
Unk Unk Unk
Polyhedral, 50-60 nm in diameter Polyhedral, 175 nm in diameter Polyhedral, 150-180 nm in diame-
Unk Unk Unk
175 Carson and Brown (unpublished) cited in 149 55 152 127
Paraphysomonas corynephora'
Cytoplasm
Unk
Unk
127
Paraphysomonas bourrellyi
Cytoplasm
Unk
Unk
127
Unk Unk
Unk Unk
Polyhedral, 22 nm in diameter Polyhedral, 22 nm in diameter
Unk Unk
82 82
Nucleus?
Unk Unk
Polyhedral, 65 nm in diameter Polyhedral, 400 nm in diameter
Unk Unk
Sirodotia tenuissimad Cryptophyceae *Cryptomonas sp.'
Nucleus and cytoplasm Cytoplasm
nutah
Chrysophycean-like'
ca'
Prymnesiophyceae (Haptophyceae) *Chrysochromulina sp. *Coccolithus huxleyi *Hymenomonas carterae Strain ln
Cytoplasm
Strain 2
ter Three polyhedral particles, 50-60 nm, 150-180 nm, and 270-300 nm in diameter Two polyhedral particles, 50-60 nm and 150-180 nm in diameter
124
Cooper and Brown (unpublished) cited in 149
Eustigmatophyceae Monodus sp.°
Nucleus?
Unk
Polyhedral, 400 nm in diameter
Unk
Huang and Hommersand (unpublished) cited in 149
Phaeophyceae *Chorda tomentosaP *Ectocarpus fasciculatisq
Cytoplasm? Nucleus?
Unk Unk
Polyhedral, 170 Polyhedral, 170
nm nm
in diameter in diameter
Unk Leads to cell lysis Infects swimming zoo-
176 4, 21
*Ectocarpus
siliculosusr
Cytoplasm
dsDNA
Polyhedral, 130
nm
in diameter
*Feldmannia
sp.S
Nucleus?
dsDNA
Polyhedral, 150
nm
in diameter
spores
Replicates only in
*Pylaiella littoralis'
*Sorocarpus uvaeformisu *Streblonema sp.v
103 51
meiotic
sporangia
Nucleus then cytoplasm?
Unk
Cytoplasm Cytoplasm
Unk Unk
Polyhedral, 130-170
nm
in diame-
Unk
83
Infectious Unk
117 71
ter
Polyhedral, 170 nm in diameter Polyhedral, 135-150 nm in diameter
Continued on following page
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
589
TABLE 1-Continued Host specieSa
Prasinophyceae *Heteromastix sp.
Location of virogenic stroma
Nucleus?
Nucleic acid
Unk
Morphological
characteristics
Polyhedral, 320 nm, with complex membranes
Infection and replication facts
Unk
KeY references e
Ott and Hommersand (un-
published) Mesostigma viridew *Micromonas pusillax
Cytoplasm
*Platymonas sp.Y *Pyramimonas orientalisz
Cytoplasm
Unk dsDNA
Polyhedral, 130 nm in diameter Polyhedral, 130-135 nm in diameter
Nucleus Nucleus
Unk Unk
Polyhedral, 51-58 nm in diameter Two polyhedral particles, 60 and 200 nm in diameter
Chlorophyceae Aulacomonas Sp.aa
Cytoplasm?
Unk
*Brachiomonas sp.
Cytoplasm?
Unk
Chlorella spp.bb
Unk
Unk
Chlorella spp.Cc Chlorella strain NC64Add
Cytoplasm
Cytoplasm
dsDNA dsDNA
Polyhedral, 200-230 nm in diameter, with tail 150-200 nm long Polyhedral, 380-400 nm in diameter; some had long tails (500 nm) Polyhedral head, 41 nm in diameter, with a 24-nm-long, 10-nmwide tail Polyhedral, 170-180 nm Polyhedral, 150-190 nm
Chlorella strain Pbiee
Cytoplasm
dsDNA
Polyhedral, 140-150 nm
Unk Virus adheres to the surface of host cells, not known if uncoating occurs at the surface or internally Unk Unk
cited in 149 97 87, 124, 168, 199
122 101
Unk
170
Unk
54
Unk
174
Unk Attaches specifically to host walls and uncoats at surface Attaches specifically to host walls and un-
96, 186, 187 63, 181, 184, 187
132, 134
coats at sur-
Chlorococcum minutumrf
Cytoplasm
dsDNA
Polyhedral, 180-220 nm in diameter, has a tail
Cylindrocapsa geminellagg
Nucleus, then cytoplasm?
dsDNA
Polyhedral, 200-230 nm in diameter with a membranous coat
Oedogonium sp.h Radiofilum transversale" Stigeoclonium farctum"
Cytoplasm
Polyhedral, 240 nm in diameter Polyhedral, 41 nm in diameter
Chloroplast?
Unk Unk Unk
Uronema gigaskk
Cytoplasm?
dsDNA
Chara corallina"'
Cytoplasm?
ssRNA
Tubular rods, 532 nm long, 18 nm wide, with basic pitch of 2.75 nm
Coleochaete scutata
Nucleus
Unk
Polyhedral, 41 nm in diameter
Nucleus
Tubular? 16.3 nm in cross-section Polyhedral, 390 nm in diameter, some have long tails (1,000 nm)
face Virus infects only zoospores, progeny viruses appear in ca. 8 h, viruses released by lysis of cell Particles may lead to cell lysis Unk Unk Unk Unk
46, 47
56, 163, 164 123 86 86
23, 29, 86
Charophyceae By artificial in38, 155 jection of virus into uninfected Chara cells; chlorosis and death in 10-12 days 86 Unk Continued on following page
590
VAN ETTEN ET AL.
MICROBIOL. REV.
a Classification from reference 141. Asterisks denote marine algae. b The intranuclear particles (termed concentrosomes) are probably not VLPs. The polyhedral particles are present only in the cytoplasm of a small number of cells. c Unk, unknown. d The virogenic stroma is prominent in the apical cell and appears to be transmitted to the daughter cells during division of the apical cell. The host cells with viral particles may be scenescent. f Cells with particles lack a nucleus. g Franca (34) reported two types of VLPs. Soyer (162) suggested that the V2 particles are tricocystoid filaments. The Vl particles do not have helical symmetry and are unlikely to be VLPs. h Two classes of VLPs were sometimes present in the same cell. 'The unidentified chrysophycean alga was an endosymbiont in the dinophycean alga Peridinium balticum. i This VLP may be responsible for rapid cell lysis when natural collections are brought to elevated temperatures of the laboratory, or in late spring when the water temperature increases. This may explain the mass-scale lysis of Dinobryon spp. in major water supplies, giving rise to subsequent blooms of other algae. k VLPs were probably present in zoospores and were not seen in vegetative cells. 'The VLPs may lyse the alga. The VLPs were seen in 75% of the algal cells after enrichment for the alga, which was followed by the disappearance of the alga. m Three size classes of VLPs were observed in the alga; the two smaller particles were occasionally present in the same cell. n Cells containing VLPs were greatly enlarged and contained proliferating chloroplasts. 0 The host could be xanthophycean or intermediate between the Xanthophyceae and Eustigmatophyceae. P VLPs found in settled zoospores of Chorda; spores with VLPs do not synthesize a cell wall and cell organelles were disrupted. e VLPs found in the plurilocular sporangia of native material. VLPs also found in developing zoospores; thus the VLPs may have been transmitted through the zoosporangium to the zoospore by subsequent cleavage. Particles are associated with tubular elements. The particles appear only in gametangia of certain isolates. They have a dsDNA of ca. 350 kbp. No nuclear membrane is present in sporangia with VLPs, but surrounding matrix appears to be nucleoplasm. They have a dsDNA of about 190 kbp. VLPs are present only in sporangia and are not seen in vegetative cells. VLPs are usually present in zoospores but are also seen in vegetative cells of the alga. VLPs are present in vegetative cells. VLPs are present in about 5% of the cells. The alga does not have a cell wall; the particles are infectious. Y Approximately 30% of the cells have particles. Both size classes of VLPs were found in the same cell. aa Aulacomonas sp. is a colorless flagellate, lacking a chloroplast. The VLPs are surrounded by an envelope about 30 nm thick. Smaller polyhedral particles (50 nm in diameter) were present in the nucleus of one cell. bb The presence of viruses in these Chlorella cells is questionable since the cultures were contaminated with bacteria and the particles may be bacteriophage. cc The host Chlorella cell is normally a symbiont of H. viridis. The VLP dsDNA is at least 250 kbp. dd The host Chlorella cell is normally a symbiont of a protozoan, P. bursaria, collected in the United States. The alga can be grown in culture, and a family of viruses which replicate in the alga can be plaque assayed. ee The host Chlorella cell is normally a symbiont of a protozoan, P. bursaria, collected in Europe. The alga can be grown in culture, and a family of viruses which replicate in the alga can be plaque assayed. ff Virus may have a retractable tail. 99 VLPs confined to single-cell germlings, and a heat shock of 40°C, 6-24 h, is necessary to induce particle formation. The heat shock, applied to the zoospores, produces germlings in which 3 to 10% of the cells are infected with VLPs. dsDNA is 275 t8 300 kbp, particles contain at least 10 proteins. hh Detected only in developing germlings. iNo pictures of these particles have been published, but they are reported to resemble the VLPs of Coleochaete spp. S Probably not a VLP. kk About 1% of the cells in the germling or young filament stage contain particles. A heat shock treatment leads to increased VLP prdduction. "The virus was the first experimental infection of a eukaryotic algal cell. Physiochemical data: s2o, = 230S; molecular weight, 3.6 x 106; base ratio of G 24.5, A 28.0, C 20.0, U 27.5; 5% RNA; protein coat molecular weight, ca. 17.5 x 103.
Symbiotic Chlorelia Species Most algae assigned to the genus Chlorella are free living in nature. However, some forms called zoochlorellae or Chlorella-like algae, live as hereditary endosymbionts wvithin freshwater and, to a lesser extent, marine animals (see, e.g., references 40, 105, 159, and 178). For convenience, we will use the term zoochlorellae in this review. Douglas and HIuss (31) have shown that the DNA from several zoochlorellae hybridize with free-living C. vulgaris DNA. Because of the similarity of zoochlorella DNA and C. vulgaris DNA and because some zoochlorellae can be cultured, two groups of investigators (31, 62) have suggested that zoochlorella symbiotic associations may have arisen relatively recently. The green coelenterate H. viridis (106) and the unicellular green protozoan P. bursaria (61) are among the most widely studied organisms containing zoochlorellae. Each gastrodermal cell of H. viridis contains 10 to 20 zoochlorellae and a single hydra contains 1.5 x 10i zoochlorellae under optimum laboratory conditions. P. bursaria typically contains several hundred zoochlorellae. Each algal cell is enclosed within an individual, hostderived vacuole, known as the perialgal vacuole (193). The algae are always located in a well-defined position in the cells of the host hydra or paramecium (159). The zoochlorellae
grow and divide in concert with the host so that the population of zoochlorellae is maintained at a constant level. When green hydras and green paramecia are placed under nongrowing conditions, algal growth also ceases. Thus the zoochlorellae and host grow coordinately (see, e.g., reference 32). Symbiotic zoochlorellae fix CO2 in litht and can transfer 30 to 40% of their total photosynthate to the host hydra or paramecium (33, 98, 131). The symbiotic relationship can also be maintained for at least a short time in dark-grown hydras (120) or paramecia (89). Symbiont-free Hydra viridis can be obtained in the laboratory by growing hydras in glycerol or in high-intensity light either alone or in the presence of a photosynthietic inhibitor such as 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) (121). These treatments produce a white, aposymbiotic hydra. The aposymbiotic hydra can be grown in culture indefinitely if appropriately fed. In contrast, most investigators have had trouble growing hydra zoochlorellae in culture. Although there is one report of the successful culturing of a hydra zoochlorella (59), others believe that this alga may have been a surface contaminant (66).. Aposymbiotic P. bursaria organisms are produced by growth in the dark at high growth rates for several months
VOL. 55, 1991
(130, 154). Paramecium zoochlorellae are apparently easier to culture than those of hydras, since they have been cultured on several occasions (see, e.g., references 75, 119, 154, 201). Cultured zoochlorellae release as much as 80% of their photosynthate to the environment (17), primarily in the form of the disaccharide maltose, whereas free-living Chlorella species do not. Maltose release is pH dependent; it is maximal at about pH 4.0 and declines with increasing pH to nearly zero at pH 7.0. The excreted maltose apparently does not arise directly from starch degradation; instead, Cernichiari et al. (17) suggested that a specific glucosyltransferase condenses glucose at the algal cell surface to produce maltose. The symbiotic relationships of hydras or paramecia and their zoochlorellae can be reestablished by coculture of the host and its symbiont. Symbionts are individually taken up into perialgal vacuoles, which' avoid fusion with lysosomes. The fate of nonsymbiotic Chlorella cells taken in by hydras or paramecia varies with the Chlorella species and host. Typically, the nonsymbiotic Chlorella cells enter food vacuoles and are digested as the vacuoles fuse with lysosomes or else the Chlorella cells fail to grow and divide (see reference 90 and references cited therein). Hereditary endosymbiotic associations are usually, but not always, host specific. For example, reassociation studies with different Chlorella species and aposymbiotic paramecia indicate that the colorless paramecium can take up not only its original symbiotic partners, but also a limited number of other Chlorella species, and can form green paramecia under special conditions. However, only the original paramecium zoochlorellae can reestablish a long-term, stable symbiosis with the protozoan (131). Paramecium zoochlorellae can also form a symbiotic association with hydras which is stable under laboratory conditions (91, 106). Kessler et al. (66) reported that a variety of free-living Chlorella strains could form a symbiotic relationship with hydras. The single characteristic- correlated with this property was the ability of the algae to grow under acidic (pH 4) conditions. This finding, if verified, suggests that specific surface recognition may not be as important in establishing symbiosis as was once thought (see, e.g., references 95 and 125). However, maintaining a stable symbiotic relationship is more complex. The ability of the alga to supply the host with carbon, in the form of maltose (159), and to utilize nitrogen and other host nutrients is probably important. Understanding symbiosis will require discovering the details of the physiological adaptations.
Chlorella Strains NC64A, Nla, Pbi, and SAG-241-80 The hosts for two groups of Chlorella viruses are the exsymbiotic Chlorella strains NC64A and Nla, isolated almost 30 years apart from P. bursaria collected in the United States, and Chlorella strains Pbi and SAG-241-80, isolated from paramecia collected in Europe. Chlorella strains NC64A and Nla have identical mitochondrial and chloroplast genomes (91), and thus the algae are probably identical. We have used NC64A and Nla interchangeably in our virus studies, and to simplify the discussion we will refer to both strains as NC64A in this review. Chlorella strain NC64A can be grown easily in the laboratory on a simple salt solution (Bold's basal medium [111]) modified by adding 0.5% sucrose and 0.1% protease peptone at 25°C in continuous light (ca. 30 microeinsteins m-2 ') with gentle shaking. Under these conditions the alga pro-
VIRUSES OF EUKARYOTIC ALGAE
591
duces four autospores about every 48 h. The algae are oval and measure about 4 by 5 ,um. They have a single cupshaped chloroplast with a prominent pyrenoid. Commercially available enzymes digest the cell walls of some free-living Chlorella species to give viable protoplasts, e.g., C. vulgaris, C. saccharophila, and C. ellipsoidea (1, 7, 220). Chlorella strain NC64A is resistant to commercial cell-wall-degrading enzymes (91). However, we have partially purified an enzyme(s) from virus lysates which degrades Chlorella strain NC64A cell walls. This enzyme(s) (called lysin) can produce protoplasts of Chlorella strain NC64A. As measured by vital stains, the protoplasts live for up to 4 weeks, but attempts to regenerate colonies from these protoplasts have so far been unsuccessful (91). Isolated Chlorella strain NC64A cell walls were analyzed for component monosaccharides (92). Glucose and rhamnose made up 51 and 16% of the total sugars, respectively, with galactose, xylose, arabinose, mannose, and glucosamine each accounting for 5 to 10% of the sugars. These monosaccharides made up about 50% (by weight) of the solubilized wall material. The nature of the remaining wall material is unknown. Chlorella strain NC64A nuclear DNA is 67% G+C (189), and mitochondrial DNA (195) and chloroplast DNA (147) are 32 and 38% G+C, respectively. The nuclear DNA contains methylated bases; SmC makes up 21% of the cytosines, and N6-methyladenine (6mA) makes up 0.6% of the adenines (189). No 4-methylcytosine (4mC) was detected in the nuclear DNA (179). A lambda genomic library of Chlorella strain NC64A (Nla) nuclear DNA has recently been prepared in Escherichia coli K803 (91). The nuclear DNA was not clonable in other E. coli strains, presumably because of its high SmC content (194). A single-copy ,-tubulin gene, which hybridized to a Chlamydomonas tubulin gene, was identified in the Chlorella strain NC64A library and sequenced. The Chlorella tubulin gene has more introns (twelve) than any other known tubulin gene and shares sequence and gene structure with both fungi and higher plants (91). Vectors to transform Chlorella strain NC64A are being constructed with the promoter region of this tubulin gene. Chlorella strain NC64A (Nla) contains a 120-kbp circular chloroplast DNA (147), which is smaller than the 175-kbp chloroplast DNA present in free-living C. ellipsoidea (217, 218). Typically, chloroplast DNAs, including those of C. ellipsoidea, contain an inverted repetitive sequence of about 20 kbp, containing a copy of the 23S, 16S, and 5S rRNA genes. However, Chlorella strain NC64A chloroplast DNA contains only a single rRNA gene. The mitochondrial DNA from Chlorella strain NC64A (Nla) is a 76-kbp circle (195). The mitochondrial DNA was physically mapped, and some of the mitochondrial genes were placed on the map. Beyond these studies, nothing is known about the molecular properties of Chlorella strain NC64A. There is even less information about the molecular properties of Chlorella strains Pbi and SAG-241-80.
DISCOVERY OF CHLORELLA VIRUSES Initial Observations A fortuitous conversation between two of us (R.H.M. and J.V.E.) about 11 years ago led to a simple experiment, which produced an exciting result. We discovered that large (185 nm in diameter) polyhedral VLPs accumulated in zoochlorellae within a few hours after they were isolated from H.
592
VAN ETTEN ET AL.
viridis (Fig. 1). Replication of these particles lysed the entire population of zoochlorellae within 24 h following isolation from hydras (96). These VLPs, which were absent from intracellular zoochlorellae, were named HVCV-1 for H. viridis Chlorella virus. HVCV-1 was isolated and partially characterized. The particles sedimented at 2,600S in sucrose density gradients and had a density of 1.295 g/ml in CsCl. HVCV-1 contained a double-stranded DNA (dsDNA) genome of at least 250 kbp and at least 19 structural proteins varying in apparent size from 10.3 to 82 kDa. The major viral protein of 46 kDa stained with Schiffs reagent, suggesting that it was a glycoprotein (186). Our initial experiments were conducted with zoochlorellae isolated from a hydra strain (Florida strain) which had been maintained in culture for over 20 years. Subsequently we examined zoochlorellae isolated from six other hydra isolates for VLPs. In addition, zoochlorellae were isolated from P. bursaria. In each case, large dsDNA containing VLPs appeared within 24 h of zoochlorella isolation (187). Restriction endonuclease analysis of VLP DNAs revealed four distinct VLPs. The relationship of the VLPs to the source of the hydra was especially interesting. A green hydra obtained from a Massachusetts lake in 1981 yielded a VLP identical to HVCV-1. Green hydras collected in 1981 from lakes in North Carolina, Nebraska, and Massachusetts yielded a second VLP, designated HVCV-2. Two green hydras collected independently in England yielded a third VLP, desig-
nated HVCV-3. The presence of identical genomes in VLPs isolated from zoochlorellae from such diverse geographic locations suggested that the VLPs had a remarkably stable relationship with their zoochlorella host. The source of the VLPs in the hydra-zoochlorellae systems is still unknown. Particles were never observed in intracellular zoochlorellae in the hydras. The particles could result from an external infection which is maintained by the natural death of hydras and release of associated zoochlorellae. Another possibility is that association with hydras suppressed viruses lysogenic with zoochlorellae. These lysogenic viruses might even code for a gene product(s) that mediates symbiosis between the zoochlorellae and the hydras. An additional possibility is that the particles replicate in hydra cells and infect the zoochlorellae during their isolation. Although we never observed VLPs in hydra cells, there is one report of VLPs in a hydra species (Hydra vulgaris) which lacks zoochlorellae (11). These polyhedral particles were smaller (70 to 75 nm) than the VLPs subsequently found in H. viridis zoochlorellae. The possibility of lysogeny in the zoochlorellae or viral replication in the hydras could be determined experimentally; however, these experiments have not been conducted. The DNA restriction fragment pattern of the VLP from the P. bursaria zoochlorellae differed from those of HVCV VLPs, and this VLP was designated PBCV-1 for P. bursaria Chlorella virus. Kawakami and Kawakami (63) had previously observed VLPs infecting zoochlorella in a P. bursaria strain isolated in Japan. Zoochlorellae growing symbiotically inside the paramecia did not contain VLPs. However, the algae became infected immediately after release from the paramecia. The VLPs observed by Kawakami and Kawakami were neither isolated nor physically characterized; however, they were similar in size and morphology to PBCV-1. In summary, VLP infections are common in zoochlorellae shortly after isolation from hydras and paramecia. This can explain the difficulty in culturing zoochlorellae from hydras.
MICROBIOL. REV.
Although the VLP association may contribute to the symbiosis, the relationship in P. bursaria is not as strong, since zoochlorellae have been cultured free of paramecia on several occasions (75, 119, 130, 154, 201). Development of a Culturable System for PBCV-1 We tested several cultured zoochlorellae, reputedly derived from invertebrate hosts, as well as two free-living Chlorella strains, to see whether they could serve as hosts for the three HVCV VLPs and for PBCV-1. Two of the zoochlorella isolates (NC64A and ATCC 30562) and, more recently, Nla, originally from United States isolates of P. bursaria, supported virus growth (181). The progeny particles were identical to PBCV-1, and they were infectious. This fulfilled Koch's postulates, and PBCV-1 is an authentic virus. More importantly, PBCV-1 formed plaques on lawns of Chlorella strain NC64A (Fig. 2), providing a sensitive bioassay for PBCV-1. None of the HVCV viruses replicated in the zoochlorellae from paramecia, and neither PBCV-1 nor HVCV replicated in any of ca. 50 additional free-living Chlorella or Chlamydomonas strains tested to date. The plaque assay for PBCV-1, together with the ability to synchronously infect the alga, allows one to study PBCV-1 by using bacteriophage technology. For example, one-step growth studies, life cycle studies, and genetic studies are feasible. The lack of a sexual stage in the host Chlorella organism and the current absence of an effective transformation system for the alga are disadvantages of the system. After developing the PBCV-1 plaque assay, we assayed freshwater samples collected throughout the United States and other parts of the world for plaque-forming viruses. As will be elaborated in the sections on properties of Chlorella strain NC64A viruses in addition to PBCV-1 and on the natural history of the Chlorella viruses, plaque-forming viruses on Chlorella strain NC64A are common in continental American, Chinese, and Japanese, but not European, fresh water. The viruses infecting Chlorella strain NC64A are referred to as NC64A viruses. Once we realized that NC64A viruses were common, we examined our original culture of paramecia for virus by the plaque assay. This culture, which had been stored at room temperature for about 18 months, still had viable green paramecia. However, no plaque-forming viruses were detected in the culture, nor could they be detected in zoochlorellae following isolation of the zoochlorellae from these paramecia (179). Consequently, the appearance of VLPs in the original, isolated paramecium zoochlorellae probably resulted from an external infection. Plaque-forming viruses which infect Chlorella strain SAG241-80 (69, 70) and Chlorella strain Pbi (referred to as Pbi viruses) (132, 134) have also been discovered in Soviet and European fresh water. Chlorella strains SAG-241-80 and Ebi are zoochlorellae originally cultured from European isolates of P. bursaria.
PROPERTIES OF CHLORELLA VIRUSES General Characteristics of PBCV-1 PBCV-1, the prototype NC64A virus, is easy to purify, and 80 to 105 A260 units (8 to 10 mg of PBCV-1) and 1.4 x 1012 to 2 x 1012 PFU of purified PBCV-1 are obtained per liter of lysate. Purified virus can be stored at 4°C for at least 1 year without detectable loss of infectivity. However, freezing inactivates PBCV-1.
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
593
B p
11
FIG. 1. Electron micrographs of the zoochlorellae present in H. viridis (A), immediately after isolation from the hydra (B), and after incubation of the isolated algae at 20°C for 6 h (C) and 20 h (D). Note the appearance of VLPs (arrow in panel C) in the algae at 6 and 20 h after isolation. Abbreviations: g, Golgi; n, nucleus; ch, chloroplast. The bar in C represents 1 ,um. Reproduced from reference 96 with permission from Academic Press, Inc.
594
MICROBIOL. REV.
VAN ETTEN ET AL.
1 week at 22°C in 5 mM EDTA or ethylene glycol-bis
lz--""ei'll-l.N I ...
il.
::::::.:---: :f:::::":"
...
f-
..
FIG. 2. Plaque assay of PBCV-1 on a lawn of Chlorella strain NC64A. The photograph was taken 4 days after the plate was prepared. However, plaques can be observed as early as 24 h after plating.
Electron micrographs indicate that PBCV-1 is a polyhedron with a multilaminate shell surrounding an electrondense core (Fig. 3). Frozen hydrated particles measure about 175 nm in diameter (Fig. 3A) (5). More recent electron micrographs indicate that PBCV-1 may have more substructure than was originally thought. For example, particles subjected to the quick-freeze, deep-etch procedure (53) contain flexible hairlike appendages (fibers) with swollen structures at the end; these appendages extend from at least some of the virus vertices (Fig. 3B). Micrographs of negatively stained particles also suggest that long fibers are attached to the outside of the capsid (small arrow in Fig. 3C). In addition, these micrographs suggest that some particles contain a distinctive 20- to 25-nm spike extending from one vertex (large arrow in Fig. 3C). The proportion of such particles suggests a unique vertex (50). DNA within the particle seems to be retracted from this vertex. The diameter of PBCV-1 is about 160 nm by comparison with lambda phage on the same grid (50). Speculation on the possible function of both the hairlike appendages and the spike structure will be mentioned in the section on virus attachment. It is obvious that additional research on the ultrastructure of PBCV-1 is required and that the particle may be more complex than a simple polyhedron. PBCV-1 sediments in sucrose density gradients at about 2,300S. The virus is disrupted on CsCl and Cs2SO4 density equilibrium gradients (187). The molecular weight of PBCV-1 has been estimated by two methods. From the DNA content (21 to 25%) and size (2.1 x 108 Da), the virion has a weight of 0.84 x 109 to 1 x 109 (184). Field flow fractionation gives a molecular weight of 1 x 109 (222). If one assumes a viral molecular weight of 1 x 109 and an extinction coefficient A 260 of 10.7 (uncorrected for light scattering), one A260 unit of PBCV-1 contains about 5.6 x 1010 virus particles (184). Experimentally we obtain 1.5 x 1010 to 3 x 1010 PFU per A260 unit of PBCV-1. Thus, 25 to 50% of the viral particles form plaques. PBCV-1 was treated with several reagents under a variety of conditions in an attempt to produce empty capsids and/or stable subviral core particles (156, 188). PBCV-1 is stable at pH 4 to 11. Virus infectivity is rapidly lost if PBCV-1 is stored above 45°C. PBCV-1 infectivity was stable for at least
(P-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA);
however, infectivity decreased slowly in 50 mM EDTA or EGTA. Virus treated with either 5 mM dithiothreitol or 5 mM dithioerythritol, but not 5 mM mercaptoethanol, rapidly lost infectivity. The dithiothreitol- or dithioerythritol-treated virus sedimented at the normal rate in sucrose gradients; however, electron micrographs of uranyl acetate-stained preparations revealed disrupted virus. To date, none of the treatments which decreased PBCV-1 infectivity have yielded either empty capsids or stable subviral core particles. Composition of PBCV-1 PBCV-1 contains about 64% protein, 21 to 25% dsDNA, and 5 to 10% lipid (156, 184). Analysis of PBCV-1 proteins by one-dimensional sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Fig. 4A) indicates more than 50 structural proteins ranging in apparent size from 10 to more than 200 kDa. Two-dimensional PAGE revealed additional proteins. However, some of the spots (especially the major capsid protein) smear on isofocusing gels, and repeated attempts to obtain "clean" two-dimensional patterns have been unsuccessful (157). One 54 kDa protein (Vp54) makes up about 40% of the total viral protein. If PBCV-1 is heated in denaturing buffer at 60°C, instead of 100°C, Vp54 migrates as a dimer with an apparent molecular weight of 104,000 (Fig. 4A, compare lanes 1 and 2). Vp54 is a glycoprotein. After deglycosylation of the protein with trifluoromethanesulfonic acid or hydrogen fluoride-pyridine, the protein migrates with an apparent size of about 49 kDa (196). The nature of the glycan linkage is unknown, although two observations suggest that it is 0 linked. Monensin, but not tunicamycin, inhibits both PBCV-1 synthesis and Vp54 synthesis (198). Incubation of Vp54 with the enzyme endo-,-N-acetylglucosaminidase, which cleaves N-linked sugars, does not affect its migration on SDS-gels. A second protein, Vp135, also stains with Schiff's reagent and is probably a glycoprotein. Vp135, Vp7l, Vp54, and Vpl4.5 are labeled by treating intact virus with 125I and are believed to be surface proteins (156). Serological experiments, described below, indicate that the glycan is on the virus surface. The resistance of PBCV-1 to several proteases is consistent with this interpretation. PBCV-1 proteins were also analyzed for myristylation and phosphorylation by infecting cells in the presence of [3H]myristic acid or 32P043- (128). Four proteins (Vp135, Vp55, Vp54, and Vp27.5) were specifically labeled by myristic acid. The myristic acid label was resistant to hydroxylamine (pH 8.5) and alkaline methanolysis, suggesting an amide linkage. Six PBCV-1 proteins (Vp6O, Vp45, Vp36, Vp29, Vp2O, and Vpl4) were labeled with 32P043- suggesting that they are phosphoproteins. The major lipids in PBCV-1 are phosphatidylcholine, phosphatidylethanolamine, and an unidentified component (156). PBCV-1 loses infectivity rapidly in chloroform and more slowly in ethyl ether or toluene; thus the lipid is required for infectivity. Sedimentation of organic solventtreated particles on sucrose gradients revealed that chloroform disrupted the virus. Ethyl ether treatment for 1 week did not alter the sedimentation properties of the virus. However, the ether-treated virus was swollen and partially disrupted after staining with uranyl acetate. PBCV-1 infectivity is unaffected by incubation with several detergents. For example, 2% Triton X-100, 2% Nonidet P-40, and 2% sodium deoxycholate have no effect on viral
* . e : . : *: : .Fse .s:i
*: A ..:. h. :.:S :.: SS: .::ee: {
i
.eSZ..Z.
*+S W: ;A: *\eS>{....;
':
:e
.: .:.
:e
..
..
..
.:
.:
.. ..
.::
.:.
..
.:
::
.: e.,
t-'.st>
.. *:
.;.
..
.:
_
..
.:
FIG. 3. Morphology of PBCV-1. (A) Frozen-hydrated particles (photograph kindly provided by Tim Baker). Note that the capsid appears as a double-layered structure in some of the particles. Bar, 200 nm. (B) PBCV-1 subjected to the quick-freeze, deep-etch procedure (53) (photograph kindly provided by John Heuser). Note the presence of flexible hairlike fibers, which appear swollen at their ends, on the surface of the particles. (C) PBCV-1 negatively stained with uranyl acetate (photograph kindly provided by Roger Hendrix). The smaller particles are lambda phage. Note that (i) long fibers are associated with the particles (small arrow), (ii) a distinctive 20- to 25-nm spike structure (large arrow) extends from one vertex of the particle, (iii) the particle appears to be packed asymmetrically, and (iv) an interior membrane appears to be located inside the capsid which extends to the unique vertex. 595
596
MICROBIOL. REV.
VAN ETTEN ET AL.
A
B 2
1 -w
BA
PST II
_MHI ID I H.
_l :.4
8268-
OR
0.
29-
V ,
_
4
.
dam "-a
a0wu
_e
17.2-
*FIG. 4. SDS-polyacrylamide gel of PBCV-1 proteins and PBCV-1 DNA restriction fragments. (A) Fluorogram of "4C-labeled PBCV-1 structural proteins following incubation at 60°C for 1 h (lane 1) or at 60°C for 55 min followed by 100°C for 5 min (lane 2). The material marked with an asterisk is presumed to be lipid. Note that the major viral protein, Vp54 (0), exists as a dimer in lane 1 and as a monomer in lane 2. Reproduced from reference 156 with permission from Academic Press, Inc. (B) PBCV-1 DNA restriction fragments: BamHIII (lane 1), HindIll (lane 2), and PstI (lane 3). Reproduced from reference 39 with permission from Martinus Nijhoff Publishers.
infectivity. However, PBCV-1 loses infectivity slowly in 2% sodium N-lauroyl sarcosine and rapidly in 2% SDS. These data, together with the ultrastructural data (i.e., an internal lipid layer is apparent in Fig. 3C), suggest that the lipid component is located inside the outer glycoprotein capsid. PBCV-1 Genome PBCV-1 contains a large dsDNA genome with a 40% G+C content (189). PBCV-1 DNA contains methylated bases. 5mC makes up 1.9% of the total cytosines, and 6mA makes up 1.5% of the total adenines. The size of the PBCV-1 genome, 333 kbp, was derived initially from summing restriction fragments. PBCV-1 DNA contains at least 41 BamHI fragments, 44 HindIIl fragments, and 50 PstI fragments (Fig. 4B) (39). Analysis of intact DNA by pulsed-field electrophoresis verified the size of PBCV-1 DNA at about 333 kbp (138). We initially reported that the PBCV-1 DNA restriction map was circular (39). This map was based on analyses of both cloned DNA and Southern transfers probed in some cases with gel-purified fragments that could not be cloned. From these experiments we concluded that PBCV-1 DNA was either a covalently closed circle, circularly permuted, or linear with terminal repetition. Subsequent experiments have established that the genome is a linear nonpermuted molecule and that the termini contain identical inverted
repeats. Furthermore, the termini have covalently closed hairpin ends (138, 167). PBCV-1 DNA terminal fragments were identified by incubating intact DNA with Bal31 exonuclease, which can initiate digestion in the single-stranded region of hairpin ends (see, e.g., references 6, 9, and 221), for various times before digestion with either HindIII, BamHI, or PstI restriction endonucleases (138). Two HindIII fragments (H4 and H17), two BamHI fragments (BlO and B15a), and two PstI fragments (P17 and P31) disappeared with increased Bal 31 exonuclease treatment, suggesting that these fragments were termini. Hybridization studies established that restriction fragments BlSa, H4, and P31 (arbitrarily called the left end) were at one terminus and that B10, H17, and P17 were at the other terminus. A revised restriction map beginning with Bl5a at zero and ending with B10 at 333 kbp is shown in Fig.
5. The terminal fragments, e.g., P31 and P17, hybridize with another, indicating terminal repetition. The termini (minus the hairpin) have recently been cloned and sequenced to within about 40 to 60 bases of the end (167). The termini contain identical inverted repeats of at least 2,185 bases, after which the sequence diverges. The inverted repeats contain two small open reading frames (ORFs) and several direct repeats. However, neither of the ORFs nor the remainder of the inverted repeats appear to be transcribed one
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
597
B31
B1 5a H4 P31
B2
I
I
I
P24
| |P14 P36a
0
|
H12a
H27b
|PS 10
B141 616 B8a HI8aI H24bI H23b I HIa P1O
20
IB2311
B12b
u
P13
B7b
I
I P9a 7080
P34b
Bib
I
H14
P4
|
110
H19 P15 I
50
Blb I P22
H3
Pi8c
I
100
Bla HI 8b
Hib
|
H29 130
P3b
P28b
B17 H13 90
B7c
|
I
H21 |
40
I H22bI
P26a 1,1 I P35 0 P38 P39
|HI8a |P25b
B32
II I
B14
H8b
P30 |P3a 30
P37
50
1 100
I
| H8c
B28
H3 P21
l'IB241
B8b
120
P32
I
P2c
140
150
B30
B1a H1b P2c
|
I
B12a
PI8b
i
150
619 |
|
|I H28
16
H9
P2a
170
160
B11
| B21|
H7
B4
H16
I
P28a
I
HUa
P33
IP34a4
P11
1;O
180
200 B25
84 HUa
|618b
-
P 1
|, I H24a| H27a
P8
200
1B26
63
| 620b
IH25 H23c
H20b
H15
PS
210
7a
||
H2 P27
220
P9b
|
P23
230
P25a
|
240
250
629 B7a H5c J P18a
B9 |
B5 Hil H
,H26| P29 PlBa
P16b
B6
|H20a
6B15b
300
320
B22 H22a
I
H5b
P16c
H17 P20
310
P7
B13
610
|H8a
P2b
H1O
H12b
P26b 280 P36b
270 | B20a
H5b
B18a H23a
P12
260
250
8B27
P17
330
333
FIG. 5. Revised PBCV-1 restriction map. The restriction sites for BamHI, HindlIl, and PstI are shown. Fragments Bl5a, H4, and P31 were placed at the left terminus, coordinate zero, and fragments B10, H17, and P17 were placed at the right terminus.
during PBCV-1 replication. The remainder of the PBCV-1 genome contains primarily unique DNA and has the potential to code for several hundred proteins (39). Serology of PBCV-1 Polyclonal antiserum prepared against intact PBCV-1 completely inhibits PBCV-1 infection by agglutinating the particles even at a dilution of 1:500. Western immunoblot analysis indicates that although the antiserum reacts with many of the virus proteins, it reacts most strongly with the major capsid protein Vp54 (197). If Vp54 is deglycosylated with trifluoromethanesulfonic acid or treated with periodic acid after transfer to nitrocellulose, the protein no longer reacts with PBCV-1 antiserum. Spontaneously derived antiserum-resistant mutants of PBCV-1 can be isolated at a frequency of about 10-6. Polyclonal antiserum was initially prepared against one of these mutants, named EPA-1. EPA-1 antiserum completely inhibits EPA-1 infection (dilution of 1:500), but has no effect (dilution of 1:10) on PBCV-1 infection (19). EPA-1 major capsid protein migrates slightly faster, with an apparent molecular mass of 53 kDa, than PBCV-1 Vp54. Like antiserum to PBCV-1, antiserum to EPA-1 reacts with the glycosylated but not the deglycosylated form of the major capsid protein on Western blot analysis. Subsequently, we isolated mutants resistant to both PBCV-1 and EPA-1 antiserum and prepared polyclonal
antiserum against some of these mutants (197). The antiserum-resistant mutants have several interesting properties. (i) Four PBCV-1 serotypes have been identified to date. All antisera reacted only with the virus which induced the antiserum. (ii) The major capsid proteins from all the mutants migrated slightly faster on acrylamide gels than did those of the strains from which they were derived. All major capsid proteins comigrated on SDS-gels after deglycosylation. (iii) Western blot analysis of the major capsid protein from the mutants, before and after deglycosylation, indicates that the serological differences among the viruses is due to a change in the carbohydrate moiety(ies) of the major capsid protein. (iv) The ratio of seven sugars (fucose, galactose, glucose, xylose, mannose, arabinose, and rhamnose) associated with the major capsid protein of PBCV-1 and the mutants varied in a predictable manner which was related to their serotype and the migration of the major capsid protein. (v) Each of the four serotypes arises from apparent single mutations in PBCV-1. This suggests that at least four virus gene products influence the PBCV-1 serotypes and that these genes are in a common pathway involved in glycosylation. Typically, carbohydrate moieties of virus glycoproteins are added in the rough endoplasmic reticulum and Golgi apparatus by host glycosyltransferases (142). Growing a virus in a different host (e.g., herpesvirus [77, 78], and influenza virus [26, 107]) can change the nature of the glycan.
598
VAN ETTEN ET AL.
Mutations that alter the polypeptide directly, e.g., changing a serine or asparagine to another amino acid (e.g., vesicular stomatitis virus [79]), can eliminate the glycan. We are presently testing the mutation possibility by cloning and sequencing the major capsid protein from PBCV-1 and some of the serological mutants. If the major capsid protein in PBCV-1 and the serotypes have identical amino acid sequences, this will confirm direct participation of viral enzymes in glycosylation.
Additional Chlorella Strain NC64A Viruses Hundreds of plaque-forming viruses, all of which infect Chlorella strain NC64A, have been isolated from fresh water collected throughout the continental United States and from China. Thirty-six of these viruses have been partially characterized and can be grouped into 16 classes (Table 2). Like PBCV-1, each of these viruses contains many structural proteins and a large dsDNA genome (at least 300 kbp) with 40% G+C, and all are chloroform sensitive. The viruses are polyhedra, with diameters of 150 to 190 nm. The DNAs of some hybridize extensively with PBCV-1 DNA, whereas others hybridize poorly (144, 227). The viruses can be distinguished from PBCV-1 and from each other by plaque size, antiserum sensitivity, DNA restriction patterns, sensitivity of the DNAs to restriction endonucleases, and the nature and abundance of methylated bases. DNA from each of the viruses contains 5mC in amounts varying from 0.12 to 47.5% of the total cytosine (Table 2). In addition, 24 of the 36 viral DNAs contain 6mA in amounts varying from 1.45 to 37% of the total adenine. The virus DNAs do not contain 4mC. Thus these viruses are a rich source of methylated DNAs. Such DNAs may be useful for studying the biological and physical properties of methylated DNAs. Growth curves revealed that viruses that form plaques similar in size to PBCV-1 (e.g., NC-1A and IL-3A) replicate at about the same rate as PBCV-1, whereas viruses that form small plaques (e.g., SC-1A, CA-4A, and NY-2A) take about two to three times longer to replicate (190). The burst size for each of the viruses is typically 200 to 350 PFU.
Viruses Infecting Chlorella Strain Pbi Plaque-forming viruses that infect an exsymbiotic Chlorella strain (Chlorella strain Pbi) of a European isolate of P. bursaria have also been isolated from European fresh water (132, 134). The Pbi viruses, like the NC64A viruses, are large polyhedra with diameters of 140 to 150 nm, are chloroform sensitive, have many structural proteins, and have large dsDNA genomes of at least 300 kbp that contain methylated bases (Table 3). However, the Pbi viruses are serologically distinct from the NC64A viruses, and their DNAs hybridize poorly with the NC64A virus DNAs. The Pbi virus DNAs have a higher G +C content (46%) than do the NC64A virus DNAs (40% G+C). Pbi viruses neither infect nor attach to Chlorella strain NC64A and the NC64A viruses neither infect nor attach to Chlorella strain Pbi (133). Additional viruses infecting Chlorella strain SAG-241-80 have been isolated and characterized from Soviet fresh water (69, 70). These viruses also infect Chlorella strain Pbi. Thus Chlorella strains Pbi and SAG-241-80 may be closely related or identicat.
MICROBIOL. REV.
REPLICATION OF PBCV-1
One-step growth experiments revealed that progeny PBCV-1 is first released about 3 to 4 h after infection and that virus release is complete by 8 to 10 h (Fig. 6A) (184). PBCV-1 release involves localized lysis of the cell wall. Thus one or more late virus gene products presumably degrade cell walls. Mechanical disruption of the cells releases infectious virus 30 to 50 min prior to spontaneous lysis. Consequently, infectious PBCV-1 is completely assembled inside the host and does not acquire lipids by budding through the host membrane. PBCV-1 also replicates in dark-grown Chlorella cells (Fig. 6B) or in light-grown cells treated with the photosynthetic inhibitor DCMU prior to virus infection. In both cases the time course of virus release is similar to that of untreated light-grown cells, and so PBCV-1 replication does not require host photosynthesis. However, the burst size, which is 200 to 350 PFU per cell in light-grown cells, is reduced about 50% in dark-grown cells. PBCV-1 infection rapidly inhibits host CO2 fixation. PBCV-1 replicates most efficiently in actively growing host cells and poorly in stationary-phase cells. Attachment of PBCV-l to Chlorella Strain NC64A PBCV-1 attaches rapidly and irreversibly to the external surface of cell walls, but not to protoplasts, of Chlorella strain NC64A with an adsorption rate of 5 x 10-9/ml/min. PBCV-1 attachment is specific since the virus attaches only to its host (93). Attachment occurs between pH 3.5 and 9 and between 5 and 25 mM MgCl2 or 5 and 100 mM NaCl (92). Each cell contains at least 5 x 104 binding sites, and Scatchard analysis indicates that 5,000 PBCV-1 particles can adsorb to each cell. Assuming that the alga is a sphere with a diameter of 5 ,Lm (94), approximately 2,500 virus particles of 175 nm would cover the surface of a single cell. Other estimates of the size of PBCV-1 have been smaller, 160 nm (50) or 139 nm (222), which means that more virus particles could fit on the surface of a single cell. From these data we concluded that (i) the entire surface of the Chlorella cell wall contains PBCV-1 receptors, (ii) individual receptor sites have similar affinities, and (iii) at most only a small percentage of the binding sites are used at any one time. The virus receptor is stable to heat and organic solvents since PBCV-1 attaches equally well to isolated walls of Chlorella strain NC64A even if the walls are first heated to 100°C or extracted with organic solvents, detergents, or high-salt solutions. The receptor is also resistant to many proteases, cellulase, and pectinase. However, exposure of the walls to HCl, pH 1, 0.5 M H2SO4, or 0.2 M NaOH for 1 h at 100°C inactivates the receptor (92). Treating walls with
the lectins wheat germ agglutinin, ricin, Lens culinaris lectin, or concanavalin A in the presence or absence of their specific hapten inhibitors also prevents virus attachment (91). This suggests that the lectins bind nonspecifically to the algal walls and that the lectin binding merely occludes viral attachment. With the exception of high concentrations (400 mM) of glucosamine and mannosamine, simple sugars such as glucose, N-acetylglucosamine, galactose, maltose, sucrose, fucose, rhamnose, arabinose, xylose, and N-acetylgalactosamine had no effect on PBCV-1 attachment (92). Attachment and infection of Chlorella strain NC64A by PBCV-1 is shown in Fig. 7A to E and in stereo views in Fig. 8. Attachment always occurs at a virus vertex. It is not known whether all virus vertices are identical and can serve as points of attachment. As mentioned above in the section
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
599
TABLE 2. Separation of Chlorella strain NC64A viruses into 16 classes' Class
1
Reaction with antibody tob: Restriction Plaque Level of methylation (mm) NY-2C Virus(m)gopMC group NY-2A 5mCd 6mAe PBCV-1 NYs-1 size
NE-8D NYb-1 CA-4B
3 3 3
Yes Yes Yes
No No No
No No No
2
AL-lA NY-2C NC-lD
3 3 3
No No No
Yes Yes Yes
3
PBCV-1 NC-lC
3 3
Yes Yes
4
CA-lA CA-2A IL-2A IL-2B IL-3A IL-3D
3 3 3 3 3 3
SC-lA SC-lB
6
7
NWf
ISV No
No
A A\
No No
A A
0.12
ND ND
Partial Partial Partial
Partial Partial Partial
A A A
0.45 0.39 0.33
ND ND ND
No No
No No
No No
C C
1.86 1.72
1.5 1.6
Yes Yes Yes Yes Yes Yes
No No No No No No
No No No No No No
No No No No No No
B B B B B B
1 1
Yes Yes
No No
No No
No No
D D
1.94 2.04
7.3 7.5
NC-lA
3
Yes
No
No
No
D
7.1
7.3
NE-8A AL-2C MA-lE NY-2F CA-iD NC-lB
3 3 3 3 3 3
Yes Yes Yes Yes Yes Yes
No No No No No No
No No No No No No
No No No No No No
E E E E E E
14.3
8.1
14.9 14.6
8.1 8.1
13.4
8.2
NYs-1
1
No
Partial
Yes
Yes
F
47.5
11.3
IL-5-2s1 AL-2A MA-iD NY-2B
1 1 1 1
No No No No
Partial Partial Partial Partial
Yes Yes Yes Yes
Yes Yes Yes Yes
G G G G
45.0 35.8 47.0 36.5
16.1 14.6 16.7 16.2
10
CA-4A
1
No
Partial
Yes
Yes
H
39.8
19.6
11
NY-2A
0.5
No
Partial
Yes
Yes
I
44.9
37.0
12
XZ-3A
1
Yes
No
No
No
J
12.8
2.2
13
SH-6A BJ-2C
1 1
Yes Yes
No No
No No
No No
K K
12.6
12.8
10.3 11.5
14
XZ-6E
1
Yes
No
No
No
E
21.2
15.2
15
XZ-4C
1
No
No
Yes
Yes
G
46.7
20.8
16
XZ-5C XZ-4A
1 1
No No
No No
Yes Yes
Yes Yes
H H
42.7 44.1
27.9 28.3
5
8 9
No
No
0-44 v rl 1.60 .
ND
10.0
ND
9.4 10.9 9.7 12.6
ND ND ND ND
a The separation is based on at least one of the following criteria: plaque size, reaction with four viral antisera, sensitivity of DNA to 13 restriction endonucleases, on percentage of 5mC and 6mA in genomic DNA. b A 5-,ug sample of purified virus was mixed with twofold serial dilutions of antiserum, and the precipitate was monitored 2 h later. Yes means that a reaction
was obtained at an antibody dilution of 1:64 or greater; Partial means a dilution of 1:8 to 1:32; and No means a dilution of 0 to 1:4. ' Viral DNAs were tested for susceptibility to the 13 restriction endonucleases BglII, EcoRI, BamHI, Sall, XhoI, Bcll, Pstl, HindIII, SstI, MboI, DpnI, MspI, and HapIl. The DNAs were grouped according to the resistance of the DNAs to these enzymes: group A, DpnI; group B, Hindlll, SstI, and Dpnl; group C, Bcll and MboI; group D, Sall, XhoI, BcIl, Pstl, and Mbol; group E, Sall, XhoI, BclI, Pstl, HindlIl, SstI, and MboI; group F, BamHI, BclI, PstI, HindIII, SstI, MboI, MspI and HpaII; group G, BamHI, Sall, XhoI, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII; group H, EcoRI, BamHI, Sail, XhoI, Bcll, PstI, HindlIl, SstI, MboI, MspI, and HpaII; group I, BglII, EcoRI, BamHI, Sall, Xhol, Bcll, PstI, HindIII, SstI, MboI, MpsI, and HpaII; group J, Bcll, HindIII, SstI, and MboI; group K, Bcll, PstI, HindIII, SstI, and MboI. d Percentage of 5mC per C plus 5mC plus deoxyuridine. e Percentage of 6mA per A plus 6mA plus deoxyinosine. f ND, not detected.
MICROBIOL. REV.
VAN ETTEN ET AL.
600
TABLE 3. Properties of five Chlorella strain Pbi viruseSa Plaque
Restriction
1 1 1 1 1
M
Virus
groupb
size (mm)
CVA-1 CVB-1 CVG-1 CVM-1
CVR-1
Level of methylation 5mCC 6mAd
NDe 17.7 ND 10.1 ND
43.1 42.7 19.2 41.9 14.2
N L F L
a None of these viruses reacted with polyclonal antisera to the Chlorella strain NC64A viruses. b Viral DNAs were tested for susceptibility to the 13 restriction endonucleases BglII, EcoRI, BamHI, Sall, XhoI, BMII, PstI, HindIII, SstI, MboI, DpnI, MspI, and HpaII. The DNAs were grouped according to the resistance of the DNAs to these enzymes: group F, BamHI, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII; group L, SalI and DpnI; group M, BamHI, Sall, Hindlll, SstI, DpnI, MspI, and HpaII; group N, EcoRI, BamHI, Sall, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII. c Percentage of 5mC per C plus 5mC plus deoxyuridine. d Percentage of 6mA per A plus 6mA plus deoxyinosine. e ND, none detected.
on general characteristics of PBCV-1, certain negatively stained preparations of PBCV-1 suggest that one vertex contains a spike structure which is consistent with a unique vertex. The stereo views (Fig. 8) indicate that the virus is attached to the wall by hairlike fibers which originate from at least some virus vertices. These interesting photographs suggest that the tips of the hairlike fibers are responsible for the initial recognition and attachment of the virus to the host. They could also be involved in orienting 4 unique virus vertex to the host surface. A scanning electron micrograph of numerous PBCV-1 particles attached to the host is shown in Fig. 9.
PBCV-1 attachment is followed by dissolution of the host wall at the point of attachment (Fig. 7B to D) and entry of the viral DNA and associated proteins into the cell, leaving an empty capsid on the host surface (Fig. 7E) (93). Since the virus also attaches to and digests Chlorella strain NC64A wall fragments which have previously been heated to 100°C, the virus carries the hydrolytic enzyme(s) (Fig. 7F). The cell-wall-degrading activity is located inside the virus particle (possibly in the spike structure), because (i) intact, protease-treated PBCV-1 particles still attach and digest a hole in the host wall and (ii) cell-wall-digesting activity can be isolated from purified, osmotically disrupted PBCV-1 particles (20). This suggests that attachment might trigger a change in the virus structure. However, attachment to and digestion of wall fragments does not lead to empty PBCV-1 capsids; thus DNA release appears to require a host function(s) (e.g., perhaps the virus internal membrane fuses with the host membrane). Attempts to release DNA from PBCV-1 attached to isolated walls by adding various agents and cell fractions have so far been unsuccessful (179).
Intracellular Site of PBCV-1 Replication The destination of the infecting DNA and the intracellular site of PBCV-1 transcription and DNA replication are unknown. PBCV-1 proteins are synthesized on cytoplasmic ribosomes and not organelle ribosomes, since cycloheximide, but not chloramphenicol, inhibits viral replication (156). PBCV-1 replication does not require host transcription, since PBCV-1 replicates in UV-irradiated cells (182). Such cells are unable to form colonies, and endogenous host RNA and DNA syntheses are reduced to background levels. However, PBCV-1 replication is slow (latent period of 16 to 20 h) and the burst size is small (2 to 10 PFU per cell) in 105 -B
A 0
, -
--
-----_
-0
/1 I
0
Ji
2
104
0
1
m I
LL I
0
-i
q
2
II
G&
IL
11
03
IIp ~0
1o2
02 100
200
300
400
TlblE, MIN.
500
600 101 0
100
200
300
400
500
600
TIME. MIN.
FIG. 6. One-step growth curves of PBCV-1 on Chlorella strain NC64A grown in the light (A) or in the dark (B). Symbols: lysis; 0, mechanically disrupted algae. Reproduced from reference 184 with permission from Academic Press, Inc.
0,
spontaneous
1~~~ ~~e>6 f
A7~~~~
A. I 1A
I..,
r
s.iF.
t... ,.,.
-jt-
0.
or
.
?b
..
Pt.i
. ;rg e
I
-.
fli , sf'.
.,
½~~~~~~~~~4-~
x
'.
-.
*~~~~~~~v I.,
D
C
,W: -s >f -I "m:o;f
1:
.L
wy
,.
:,
*< 9 'i ;_ q
rI
FIG. 7. Infection of Chiorella strain NC64A by PBCV-i. (A) Viral particle in close proximity to the alga. (B and C) Attachment of PBC Vto the algal wall and digestion of the wall at the point of attachment. (D) Viral DNA beginning to enter the host. (E) An empty viral capsid remaining on the surface of the host. (F) PBC V-i attachment and dissolution of a Chiorella cell wall fragment. Note that (i) viral attachment always occurs on the external side of the wall (i.e., the internal side of the wall curls inward) and (ii) DNA is not released from viral particles attached to wall fragments. The size markers in panels E and F represent 100 nm and 200 nm, respectively. Reproduced from reference 93 with permission from Academic Press, Inc.
601
602
VAN ETTEN ET AL.
MICROBIOL. REV.
FIG. 8. Stereo views of PBCV-i attached to Chlorella strain NC64A. The cells were quick-frozen and deep-etched. (A) Side view of the particles attached to the cell wall; (B) top view of the particles attached to the cell wall. Note that the particles are attached to the wall via hairlike fibers, which appear to originate from the vertices of the virus. Photographs kindly provided by John Heuser.
UV-treated cells. PBCV-1 replication also does not require labile host factors, since the virus replicated in UV-irradiated cells that had been incubated in the dark for up to 8 h before infection. The host cells change cytologically beginning about 1 to 2 h after infection (94). In the nucleus, the nucleolus disintegrates and chromatin heterochromicity increases. The nucleus, mitochondria, and Golgi apparatus become appressed to the chloroplast, leaving one or more finely granulated electron-translucent areas in the cytoplasm (labeled V in Fig. 1OA to D). Since PBCV-1 ultimately forms in these areas, we have designated these areas virus assembly centers. Early stages of PBCV-1 morphogenesis in the virus assem-
bly centers are seen in a few cells by 1 h postinfection (p.i.) and are common by 2 h p.i. (Fig. 10 A and B). Short membranous structures, which are vesicular or angular in shape (arrow in Fig. 10 B and C), first appear in the virus assembly centers. These membranes appear to function as templates for capsid assembly. More dense material (presumably protein) appears on the exterior surface of these membranes, giving rise to angular configurations (Fig. 1OA and B). These developing capsids are bounded by a distinct electron-dense layer; a thicker translucent region is visible directly underneath this dense outer layer (Fig. lOC and D). Capsids are usually formed at the periphery of the virus assembly centers, producing the rosette pattern seen in Fig.
VIRUSES OF EUKARYOTIC ALGAE
VOL. 55, 1991
603
FIG. 8-Continued.
lOB. Apparent complete capsids are assembled prior to DNA accumulation. A ring of virus particles surrounding a virus assembly center, which are in various stages of packaging DNA, are seen in Fig. lOD. These observations suggest that PBCV-1 probably packages DNA through an opening in the capsid. By 4 to 5 h p.i. the cells contain numerous filled virus particles, which are distributed throughout the cytoplasm. Sometimes a nuclear membrane is still visible at late stages of infection (Fig. 10E). Mitochondria and the chloroplast disintegrate in late stages of PBCV-1 replication. So far there is no evidence for nuclear or organellar roles in viral replication. If the nucleus is not involved, transcriptionassociated enzymes, such as RNA polymerase(s) and pro-
cessing enzymes, must be imported into the cell with the infecting DNA. PBCV-1 has not been effectively assayed for these enzymes. We have been unsuccessful in brief attempts to detect RNA polymerase activity either in intact PBCV-1 or in PBCV-1 that had been disrupted osmotically or by freezethawing (15). Although PBCV-1 may not contain a functional RNA polymerase, we suspect that our failure may reflect unsatisfactory disruption methods. We predict that PBCV-1 imports at least some of the enzymes involved in its replication. Since PBCV-1 is assembled in specific regions in the cytoplasm, the cytoskeleton might be expected to actively target virus proteins, including the glycoproteins, to the
604
VAN ETTEN ET AL.
MICROBIOL. REV.
Although recycling from host DNA may supply some of these intermediates, it can supply only 10 to 25% of the requirement. Consequently, virus-infected cells may be an excellent model system for studying nucleotide metabolism. One obvious test for recycling is to label host DNA prior to infection and determine whether the label accumulates in virus DNA. Unfortunately, Chlorella strain NC64A lacks thymidine kinase (180) and so host DNA cannot be labeled
FIG. 9. Scanning electron micrograph of PBCV-1 particles attached to Chlorella strain NC64A (photograph kindly provided by Kit Lee). Bar, 1 plm.
assembly center. However, several cytoskeleton-disrupting agents, which inhibit either tubulin or actin functions, had no effect on PBCV-1 replication even at concentrations much higher than those required to inhibit growth of the host Chlorella cell (112). Effect of PBCV-1 Infection on Host DNA, RNA, and Protein Syntheses
PBCV-1 infection dramatically changes host macromolecular syntheses. The following changes occur in DNA metabolism (180). (i) Virus infection almost immediately inhibits host DNA synthesis. (ii) PBCV-1 DNA synthesis begins 50 to 60 min after infection and requires de novo synthesis of a protein(s). This implies that PBCV-1 directs the synthesis of at least some of its own DNA synthetic machinery. New DNA polymerase activities appear in infected cells (139, 140). (iii) PBCV-1 infection leads to the degradation of host chloroplast and nuclear DNAs within 1 to 2 h p.i. (Fig. 11). Presumably a virus-encoded protein(s) mediates this response since cycloheximide added at the time of infection prevents host DNA degradation. Host nuclear and chloroplast DNA levels also decrease, albeit more slowly, if UV-inactivated algae are inoculated with PBCV-1 (209). (iv) The total DNA level in the host cell increases 3- to 10-fold by 5 h after PBCV-1 infection. Thus PBCV-1 replication requires a large increase in deoxynucleotide intermediates.
specifically. PBCV-1 infection also alters RNA metabolism (145). (i) Viral infection rapidly inhibits host RNA synthesis. (ii) Chloroplast rRNAs, but not cytoplasmic rRNAs, are degraded beginning about 30 min p.i. Presumably a virusencoded protein(s) contributes to this process, since cycloheximide added at the time of PBCV-1 infection prevents chloroplast rRNA degradation. (iii) PBCV-1 transcription is programmed, and early transcripts appear within 5 to 10 min after infection. (iv) Some, but not all, early viral transcripts are synthesized in the absence of de novo protein synthesis. The synthesis of later transcripts requires translation of an early gene product(s). (v) Late viral transcription begins about 1 h p.i., at the same time viral DNA synthesis begins (Fig. 12). (vi) A few early transcripts continue to be synthesized after virus DNA synthesis begins (45). Little is known about protein synthesis following virus infection. PBCV-1 infection rapidly inhibits host protein synthesis by at least 75%. We have briefly investigated virus protein synthesis in PBCV-1-infected cells by examining proteins on polyacrylamide gels (215). These experiments were less than satisfactory because (i) the synthesis of some host proteins continued (at least briefly) after infection and (ii) the specific labeling of viral proteins was low, even after long labeling periods (more than 1 h) with high-specificactivity amino acids. This latter finding suggests that PBCV-1 infection may expand amino acid pools. The increase could reflect recycling from degraded host protein or from increased amino acid synthesis. A complicating factor is that PBCV-1 has the potential to code for several hundred proteins, which would require two-dimensional gels for adequate resolution. TRANSCRIPTION OF PBCV-1 As mentioned in the preceding section, PBCV-1 transcription can be conveniently divided into early and late stages. The junction between these stages is about 1 h p.i., which coincides with the start of virus DNA synthesis. Interestingly, the incorporation of [3H]adenine into polyadenylatecontaining RNA also changes abruptly at about 1 h after PBCV-1 infection. Polyadenylate segments are virtually absent from late transcripts, whereas at least some early virus transcripts are probably polyadenylated (190). RNA isolated at various times after PBCV-1 infection was hybridized with each of four cloned fragments of PBCV-1 DNA (Fig. 12) (145). Both early and late genes are dispersed within the PBCV-1 genome, since each of the four cloned fragments hybridized to both early and late transcripts.
FIG. 10. Chlorella cells at 2 h (A to C), 5 h (D), and 6 h (E) after PBCV-1 infection. Parts of newly forming capsids (arrow in panel A) were located in defined areas in the cytoplasm (labeled v for virus assembly center in panels A to D) by 2 h p.i. Membranous materials which assume vesicular or angular shapes were usually seen in the virus assembly centers (arrow in panels B and C). The newly forming capsids can have a rosette appearance, which surrounds the virus assembly center (panel B). By 5 h p.i. (panel D) the cells contained a mixture of filled and partially filled capsids, empty capsids, and incomplete capsids. By 6 h p.i. (panel E), the cells contained many filled capsids which were distributed throughout the cytoplasm. Occasionally, cells contained aberrant virus capsids (F). Bars: 500 nm (panels A to D and F) and 1,000 nm (panel E). Reproduced from reference 94 with permission from Academic Press, Inc.
VOL. 55, 1991
~;Se$,g.A wsja35ve~ ~
VIRUSES OF EUKARYOTIC ALGAE
,,, ~ ~~~~~~~~~~~~~~~i.
i ~~~~~ N
;
5 a fi
X ;
WS2Atr 6
&
4~~~~~~~~~~~~~~~~~~~~~~1
I'
* r ve
.,... , ; ;g ,.,. ,kx !'
. , ,,:VA 3; $~I
..'r
, . . .4
AUZA
8K,K-
* A i ; ' . ?
i * '
TV;
ov
|:s;: ~~~'
eAW ^
..
;
* tj sw !Sg O
y S * * * ' + t s L " iryo4~~ ~ ~ ~ ~ ~ ~ N
i
4
w < ? >rRvntf > N.
f .Ssrv
1pm
b
V /
PIPI.e 1w,
.f
i
"Ok
C
-
0W-5).5um
_
0
d ~~~~~~~~~~~~~~~~~~~~~~~I _
_-
MICROBIOL. REV.
0. -- m
FIG. 15. Micrographs of the VLP from U. gigas. (a) Thin section of an infected cell containing hexagonal and pentagonal particles with two internal components in some particles. (b) Tailed and tailless VLPs negatively stained in 2%b phosphotungstic acid. (c) Tailed particle with a hexagonal capsid. Note the characteristic swelling in the middle of the tail. (d) Tailed particle with a swollen spherical capsid. Reproduced from reference 29 with permission from Academic Press, Inc.
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
615
nia VLPs. The Ectocarpus particles are released into the medium when host cells lyse, and they can infect healthy zoospores. The infected zoospores become paralyzed and settle to the bottom of the container, where they develop into plants that produce viruses in their sporangia. Expression of the Ectocarpus virus is temperature sensitive and is linked to differentiation of reproductive cells of the host. Notably, both research groups have preliminary evidence suggesting that the Feldmannia and Ectocarpus virus DNAs are integrated into their host DNAs (51, 104).
CONCLUDING COMMENTS
FIG. 16. VLPs near and attached to the surface of the marine nanoflagellate M. pusilla 15 min after inoculation. Unlike most eukaryotic algae, M. pusilla lacks a cell wall. Reproduced from reference 87 with permission from Macmillan Magazines Ltd.
sporangia (52), suggesting that VLP replication might be triggered by meiosis. Muller et al. (103) found viruses in Ectocarpus siliculosus, a related ectocarpoid species also isolated near the coast of New Zealand, which contained particles about 130 nm in diameter. These particles were located exclusively in the gametangia. The particles contain a 350-kbp linear dsDNA, which is about twice the size of the DNA from the Feldman-
Algal viruses, especially the Chlorella viruses, are interesting for several reasons and deserve investigation. The Chlorella viruses are examples of a new virus-host relationship, and their bacteriophagelike biology makes them excellent models for studying gene regulation and expression in photosynthetic eukaryotes. The inability to perform host genetics and to introduce foreign genes into the viruses remains a disadvantage. The genomes of these viruses are potential sources of control elements such as origin(s) of replication or promoters for plant genetic engineering. In fact, the upstream region of one of the Chlorella virus DNA methyltransferase genes (M.CviBIII) functions well as a promoter in both Agrobacterium tumefaciens and higher plants (100). Methylation levels of the Chlorella viral DNAs vary considerably; thus, some of these DNAs may be useful for molecular and biological studies on the effect of methylation on DNA structure and function. For example, it would be interesting to determine whether DNA-binding proteins
FIG. 17. Virus particles in Ectocarpus siliculosus (photograph kindly provided by Eric Henry). The particles are present only in the
sporangia.
616
VAN ETTEN ET AL.
of the viruses are adapted to levels of methylation. The Chlorella viruses contain several genes of commercial interest such as DNA methyltransferases and DNA site-specific endonucleases, some of which have unique specificities. In addition, the viruses contain host-wall-degrading enzymes which could potentially digest plant cell walls. Finally, preliminary results suggest that the viruses may contain glycosyltransferase genes. The common association of VLPs with zoochlorellae isolated from hydras and paramecia suggests that these particles may influence symbiosis. Viruses that infect Chlorella strain NC64A are common in fresh water; however, nothing is known about their potential ecological impact on the algal community. Although investigators are just beginning to look for viruses of other eukaryotic algae, it is already obvious that viruses are a major factor in marine algal ecology (153, 169). Besides limiting algal growth, the viruses could potentially mediate genetic exchange among native algae. Pearson and Norris (122) and Hoffman and Stanker (56) have also suggested that algae could serve as reservoirs or vectors for transmitting viruses to other forms of aquatic life, including animals. In this regard the Chlorella viruses have some similarities to important disease viruses such as the poxviruses and ASFV. Finally, at least some of these viruses, such as those infecting brown algae, appear to be expressed only in meiotic cells, suggesting highly specific coordination between the virus and the host life cycle. We encourage more investigators to work on these virus systems. ACKNOWLEDGMENTS We (J.V.E. and R.H.M.) thank the many technicians, undergraduate students, graduate students, postdoctoral associates, and visiting scientists who have worked in our laboratories over the past 10 years on the Chlorella viruses. In addition, many people at other institutions have graciously provided information and help on specific aspects of the viruses, as well as providing some of the photographs used in this paper. Special thanks go to Dwight Burbank, who has worked with the viruses since their discovery, and Kit Lee, who has taken many of the electron micrographs of the Chlorella viruses. We also acknowledge the many stimulating discussions with Myron Brakke over the years. A special acknowledgment goes to Malcolm Brown, who first pointed out the virus particles in one of our (R.H.M.) original electron micrographs. Finally, we thank Myron Brakke, Mike Nelson, Roy French, Reingard Grabherr, Yanping Zhang, Mike Groves, Eric Henry, Kyle Hoogland, and Jim Rosowski for reading and making many helpful suggestions on various aspects of the manuscript. Research from our laboratories has been supported by grants from the Department of Energy (J.V.E.), the National Institutes of Health (J.V.E.), the National Science Foundation (R.H.M.), the U.S. Department of Agriculture (R.H.M.), and the Solar Energy Research Institute (R.H.M.).
REFERENCES 1. Aach, H. G., S. Bortsch, and V. Feyen. 1978. Studies on Chlorella protoplasts: Demonstration of the protoplastic nature and the regeneration of the cell wall. Planta 139:257-260. 2. Adams, M. H. 1959. Bacteriophages. Interscience Publications, Inc., New York. 3. Atkinson, A. W., B. E. S. Gunning, and P. C. L. John. 1972. Sporopollenin in the cell wall of Chlorella and other algae: ultrastructure, chemistry and incorporation of "4C-acetate, studied in synchronous cultures. Planta 107:1-32. 4. Baker, J. R. J., and L. V. Evans. 1973. The ship fouling alga Ectocarpus. I. Ultrastructure and cytochemistry of plurilocular reproductive stages. Protoplasma 77:1-13. 5. Baker, T. S. Personal communication. 6. Barbour, A. G., and C. F. Garon. 1987. Linear plasmids of the bacterium Borrelia burgdorferi have covalently closed
MICROBIOL. REV.
ends. Science 237:409-411. 7. Berliner, M. D. 1977. Protoplast induction in Chlorella vulgaris. Plant Sci. Lett. 9:201-204. 8. Bestor, T., A. Laudano, R. Mattaliano, and V. Ingram. 1988. Cloning and sequencing of a cDNA encoding DNA methyltransferase of mouse cells: the carboxylterminal domain of the mammalian enzymes is related to bacterial restriction methyltransferases. J. Mol. Biol. 203:971-983. 9. Blackburn, E. H., and P. B. Challoner. 1984. Identification of a telomeric DNA sequence in Trypanosoma brucei. Cell 36:447457. 10. Blasco, R., I. D. L. Vega, F. Almazan, M. Aguero, and E. Vinuela. 1989. Genetic variation of African swine fever virus: variable regions near the ends of the viral DNA. Virology 173:251-257. 11. Bonnefoy, A. M., X. Kolenkine, and C. Vago. 1972. Particules d'allure virale chez les Hydres. C.R. Acad. Sci. (Paris) 275: 2163-2165. 12. Borowitzka, M. A., and L. J. Borowitzka (ed.). 1988. Microalgal biotechnology. Cambridge University Press, Cambridge. 13. Brooks, J., and G. Shaw. 1978. Sporopollenin: a review of its chemistry, palaeochemistry, and geochemistry. Grana 17:9197. 14. Brunt, A. A., and K. E. Richards. 1989. Biology and molecular biology of the furoviruses. Adv. Virus Res. 36:1-32. 15. Burbank, D. E., R. Grabherr, and J. L. Van Etten. Unpublished results. 16. Burbank, D. E., S. L. Shields, A. M. Schuster, and J. L. Van Etten. 1990. 5-Azacytidine resistant mutants of Chlorella virus IL-3A. Virology 176:311-315. 17. Cernichiari, E., L. Muscatine, and D. C. Smith. 1969. Maltose excretion by the symbiotic algae of Hydra viridis. Proc. R. Soc. London Ser. B 173:557-576. 18. Chapman, R. L., and N. J. Lang. 1973. Virus-like particles and nuclear inclusions in the red alga Porphyridium purpureum (Bory) Drew et Ross. J. Phycol. 9:117-122. 19. Chase, T. E., J. A. Nelson, D. E. Burbank, and J. L. Van Etten. 1989. Mutual exclusion occurs in a Chlorella-like green alga inoculated with two viruses. J. Gen. Virol. 70:1829-1836. 20. Ciuffetti, L., Q. Que, J. L. Van Etten, and R. H. Meints. Unpublished results. 21. Clitheroe, S. B., and L. V. Evans. 1974. Viruslike particles in the brown alga Ectocarpus. J. Ultrastruct. Res. 49:211-217. 22. Cock, J. M., L. L. Roof, N. F. Bascomb, C. W. Gehrke, K. C. Kuo, and R. R. Schmidt. 1990. Restriction enzyme analysis and cloning of high molecular weight genomic DNA isolated from Chlorella sorokiniana (Chlorophyta). J. Phycol, 26:361-367. 23. Cole, A., J. A. Dodds, and R. I. Hamilton. 1980. Purification and some properties of a double-stranded DNA containing virus-like particle from Uronema gigas, a filamentous eucaryotic green alga. Virology 100:166-174. 24. Danko, S., and J. L. Van Etten. Unpublished results. 25. DaSilva, E. J., and H. G. Gyllenberg. 1972. A taxonomic treatment of the genus Chlorella by the technique of continuous classification. Arch. Mikrobiol. 87:99-117. 26. Deom, C. M., and I. T. Schulze. 1985. Oligosaccharide composition of an influenza virus hemagglutinin with host-determined binding properties. J. Biol. Chem. 260:14771-14774. 27. Dodds, J. A. 1979. Viruses of marine algae. Experientia 35: 440 442. 28. Dodds, J. A. 1983. New viruses of eukaryotic algae and protozoa, p. 177-188. In R. E. F. Matthews (ed.), A critical appraisal of viral taxonomy. CRC Press, Inc., Boca Raton, Fla. 29. Dodds, J. A., and A. Cole. 1980. Microscopy and biology of Uronema gigas, a filamentous eukaryotic green alga, and its associated tailed virus-like particle. Virology 100:156-165. 30. Doermann, A. H. 1983. Introduction to the early years of bacteriophage T4, p. 1-7. In C. K. Mathews, E. M. Kutter, G. Mosig, and P. B. Berget (ed.), Bacteriophage T4. American Society for Microbiology, Washington, D.C. 31. Douglas, A. E., and V. A. R. Huss. 1986. On the characteristics and taxonomic position of symbiotic Chlorella. Arch. Microbiol. 145:80-84.
VOL. 55, 1991
32. Dunn, K. 1987. Growth of endosymbiotic algae in the green hydra, Hydra viridissima. J. Cell Sci. 88:571-578. 33. Eisenstadt, E. 1971. Transfer of photosynthetic products from symbiotic algae to animal tissue in Chlorohydra viridissima, p. 202-208. In H. M. Lenhoff, L. Muscatine, and L. V. Davis (ed.), Experimental coelenterate biology. University of Hawaii Press, Honolulu. 34. Franca, S. 1976. On the presence of virus-like particles in the dinoflagellate Gyrodinium resplendens (Hulbert). Protistologica 12:425-430. 35. Garon, C. F., E. Barbosa, and B. Moss. 1978. Visualization of an inverted terminal repetition in vaccinia virus DNA. Proc. Natl. Acad. Sci. USA 75:4863-4867. 36. Geshelin, P., and K. I. Berns. 1974. Characterization and localization of the naturally occurring cross-links in vaccinia virus DNA. J. Mol. Biol. 88:785-796. 37. Gibbs, A. Personal communication. 38. Gibbs, A., A. H. Skotnicki, J. E. Gardiner, E. S. Walker, and M. Hollings. 1975. A tobamovirus of a green alga. Virology 64:571-574. 39. Girton, L. E., and J. L. Van Etten. 1987. Restriction site map of the Chlorella virus PBCV-1 genome. Plant Mol. Biol. 9:245-257. 40. Goff, L. J. (ed.). 1983. Algal symbiosis: a continuum of interaction strategies. Cambridge University Press, Cambridge. 41. Gonzalez, A., A. Talavera, J. M. Almendral, and E. Vinuela. 1986. Hairpin loop structure of African swine fever virus DNA. Nucleic Acids Res. 14:6835-6844. 42. Goohra, R., and A. Granoff. 1979. Icosahedral cytoplasmic deoxyriboviruses, p. 347-399. In H. Fraenkel-Conrat and R. R. Wagner (ed.), Comprehensive virology, vol. 14. Plenum Press, New York. 43. Goorha, R., and K. G. Murti. 1982. The genome of frog virus 3, an animal DNA virus, is circularly permuted and terminally redundant. Proc. Natl. Acad. Sci. USA 79:248-252. 44. Goorha, R., D. B. Willis, and A. Granoff. 1977. Macromolecular synthesis in cells infected by frog virus 3. VI. Frog virus 3 replication is dependent on the cell nucleus. J. Virol. 21:802805. 45. Graves, M., and R. H. Meints. Unpublished results. 46. Gromov, B. V., and K. A. Mamkaeva. 1979. Phage-form infectious virus of the green alga Chlorococcum. Bull. Acad. Sci. USSR Biol. N2:181-186. 47. Gromov, B. V., and K. A. Mamkaeva. 1981. A virus infection in the synchronized population of the Chlorococcum minutum zoospores. Arch. Hydrobiol. Suppl. 60:252-259. 48. Hayes, W. 1968. The genetics of bacteria and their viruses, 2nd ed. John Wiley and Sons, Inc., New York. 49. Hellmann, V., and E. Kessler. 1974. Physiologische und biochemische Beitrage zur Taxonomie der gattung Chlorella. VIII. Die Basenzusammensetzung der DNS. Arch. Microbiol. 95:311-318. 50. Hendrix, R. Personal communication. 51. Henry, E., and R. H. Meints. Unpublished results 52. Henry, E. C., S. Lanka, and R. H. Meints. 1989. A stable virus infection in a brown alga. Abstr. XIII Int. Seaweed Symp., p. A98. 53. Heuser, J. 1980. Three-dimensional visualization of coated
vesicle formation in fibroblasts. J. Cell Biol. 84:560-583. 54. Hoffman, L. Personal communication. 55. Hoffman, L. R. 1978. Virus-like particles in Hydrurus (Chrysophyceae). J. Phycol. 14:110-114. 56. Hoffman, L. R., and L. H. Stanker. 1976. Virus-like particles in the green alga Cylindrocapsa. Can. J. Bot. 54:2827-2841. 57. Houts, G. E., M. Gravell, and A. Granoff. 1974. Electron microscopic observations on early events of frog virus 3 replication. Virology 58:589-594. 58. Huss, V. A. R., G. Huss, and E. Kessler. 1989. Deoxyribonucleic acid reassociation and inter species relationships of the genus Chlorella (Chlorophyceae). Plant Syst. Evol. 168:71-82. 59. Jolley, E., and D. C. Smith. 1978. The green hydra symbiosis. I. Isolation, culture, and characteristics of the Chlorella sym-
VIRUSES OF EUKARYOTIC ALGAE
617
biont of European Hydra viridis. New Phytol. 81:637-645. 60. Jones, P. A. 1984. Gene activation by 5-azacytidine, p. 165-187. In A. Razin, H. Cedar, and A. D. Riggs (ed.), DNA methylation: biochemistry and biological significance. Springer-Verlag, KG, Berlin. 61. Karakashian, M. W. 1975. Symbiosis in Paramecium bursaria. Symp. Soc. Exp. Biol. 29:145-173. 62. Karakashian, S. J., and M. W. Karakashian. 1965. Evolution and symbiosis in the genus Chlorella and related algae. Evolution 19:368-377. 63. Kawakami, H., and N. Kawakami. 1978. Behavior of a virus in a symbiotic system, Paramecium bursaria-zoochlorella. J. Protozool. 25:217-225. 64. Kelly, D. C. 1975. Frog virus 3 replication: electron microscope observations on the sequence of infection in chick embryo fibroblasts. J. Gen. Virol. 26:71-86. 65. Kessler, E. 1982. Chemotaxonomy in the Chlorococcales. Prog. Phycol. Res. 1:111-135. 66. Kessler, E., V. A. R. Huss, and M. Rahat. 1988. Speciesspecific ability of Chlorella strains (Chlorophyceae) to form stable symbioses with Hydra viridis. Plant Syst. Evol. 160:241246. 67. Klimasauskas, S., A. Timinskas, S. Menkevicius, D. Butkiene, V. Butkus, and A. Janulaitis. 1989. Sequence motifs characteristic of DNA [cytosine-N4] methyltransferases: similarity to adenine and cytosine-C5 DNA-methylases. Nucleic Acids Res. 17:9823-9832. 68. Koltin, Y., and M. J. Leibowitz (ed.). 1988. Viruses of fungi and simple eukaryotes. Marcel Dekker, Inc., New York. 69. Kvitko, K., and B. V. Gromov. 1984. New findings of a titratable infectious zoochlorella virus. Dokl. Akad. Nauk. SSSR 279:998-999. 70. Kvitko, K. V., I. I. Skoblo, E. S. Kraeva, E. V. Skovorodina, and T. A. Bulkina. 1985. Specificity of host range of zoochlorella viruses from Paramecium bursaria. All Union Meet. Microbiol. Soc. USSR 5:17. 71. LaClaire, J. W., and J. A. West. 1977. Virus-like particles in the brown alga Streblonema. Protoplasma 93:127-130. 71a.Lauster, R. 1989. Evolution of type II DNA methyltransferases: a gene duplication model. J. Mol. Biol. 206:313321. 72. Lauster, R., T. A. Trautner, and M. Noyer-Weidner. 1989. Cytosine-specific type II DNA methyltransferases: a conserved enzyme core with variable target-recognizing domains. J. Mol. Biol. 206:305-312. 73. Lee, R. E. 1971. Systemic viral material in the cells of the freshwater red alga Sirodotia tenuissima (Holden) Skuja. J. Cell Sci. 8:623-631. 74. Lemke, P. A. 1976. Viruses of eucaryotic microorganisms. Annu. Rev. Microbiol. 30:105-145. 75. Loefer, J. B. 1936. Isolation and growth characteristics of the zoochlorella of Paramecium bursaria. Am. Nat. 70:184-188. 76. Loos, E., and D. Meindl. 1982. Composition of the cell wall of Chlorella fusca. Planta 156:270-273. 77. Lundstrom, M., S. Jeansson, and S. Olofsson. 1987. Host cell-induced differences in the 0-glycosylation of herpes simplex virus gC-1. II. Demonstration of cell-specific galactosyltransferase essential for formation of 0-linked oligosaccharides. Virology 161:395-402. 78. Lundstrom, M., S. Olofsson, S. Jeansson, E. Lycke, R. Datema, and J. E. Mansson. 1987. Host cell-induced differences in 0-glycosylation of herpes simplex virus gC-1. I. Structures of nonsialylated HPA- and PNA-binding carbohydrates. Virology 161:385-394. 79. Luo, L., Y. Li, R. M. Snyder, and R. R. Wagner. 1990. Spontaneous mutations leading to antigenic variations in the glycoproteins of vesicular stomatitis virus field isolates. Virology 174:70-78. 80. Luria, S. E., J. E. Darnell, D. Baltimore, and A. Campbell. 1978. General virology. John Wiley & Sons, Inc., New York 81. Mann, M. B., R. N. Rao, and H. 0. Smith. 1978. Cloning of restriction and modification genes in E. coli: the HhaII system from Haemophilus haemolyticus. Gene 3:97-112.
618
MICROBIOL. REV.
VAN ETTEN ET AL.
82. Manton, I., and B. S. C. Leadbeater. 1974. Fine-structural observations on six species of Chrysochromulina from wild Danish marine nanoplankton, including a discription of C. campanulifera sp. nov. and a preliminary summary of the nanoplankton as a whole. Dan. Vidensk. Sel. Biol. Skr. 20:1-26. 83. Markey, D. R. 1974. A possible virus infection in the brown alga Pylaiella littoralis. Protoplasma 80:223-232. 84. Martin, E. L., and R. L. Benson. 1982. Algal viruses, pathogenic bacteria and fungi: introduction and bibliography, p. 793-798. In J. R. Rosowski and B. C. Parker (ed.), Selected papers in phycology II. Phycological Society of America Inc., Lawrence, Kans. 85. Matthews, R. E. F. 1982. Classification and nomenclature of viruses. Intervirology 17:1-199. 86. Mattox, K. R., K. D. Stewart, and G. L. Floyd. 1972. Probable virus infections in four genera of green algae. Can. J. Microbiol. 18:1620-1621. 87. Mayer, J. A., and F. J. R. Taylor. 1979. A virus which lyses the marine nanoflagellate Micromonas pusilla. Nature (London)
281:299-301. 88. McAuley, P. J., and L. Muscatine. 1986. The cell cycle of symbiotic Chlorella. IV. DNA content of algae slowly increases during host starvation of green hydra. J. Cell Sci. 85:73-84. 89. Meier, R., W. Reisser, and W. Wiessner. 1980. Cytological studies on the endosymbiotic unit of Paramecium bursaria Ehbg. and Chlorella spec. II. The regulation of the endosymbiotic algal population as influenced by nutritional condition of the symbiotic partners. Arch. Protistenkd. 123:333-341. 90. Meier, R., and W. Wiessner. 1989. Infection of algae-free Paramecium bursaria with symbiotic Chlorella sp. isolated from green paramecium. II. A timed study. J. Cell Sci. 95:571-579. 91. Meints, R. H. Unpublished results. 92. Meints, R. H., D. E. Burbank, J. L. Van Etten, and D. T. A. Lamport. 1988. Properties of the Chlorella receptor for the virus PBCV-1. Virology 164:15-21. 93. Meints, R. H., K. Lee, D. E. Burbank, and J. L. Van Etten. 1984. Infection of a Chlorella-like alga with the virus, PBCV-1: ultrastructural studies. Virology 138:341-346. 94. Meints, R. H., K. Lee, and J. L. Van Etten. 1986. Assembly site of the virus PBCV-1 in a Chlorella-like green alga: ultrastructural studies. Virology 154:240-245. 95. Meints, R. H., and R. L. Pardy. 1980. Quantitative demonstration of cell surface involvement in a plant-animal symbiosis: lectin inhibition of reassociation. J. Cell Sci. 43:239-251. 96. Meints, R. H., J. L. Van Etten, D. Kuczmarski, K. Lee, and B. Ang. 1981. Viral infection of the symbiotic Chlorella-like alga present in Hydra viridis. Virology 113:698-703. 97. Melkonian, M. 1982. Virus-like particles in the scaly green flagellate Mesostigma viride. Br. Phycol. J. 17:63-68. 98. Mews, L. K. 1980. The green hydra symbiosis. III. The biotrophic transport of carbohydrate from alga to animal. Proc. R. Soc. London Ser. B 209:377-401. 99. Mindich, L., and D. Bamford. 1988. Lipid-containing bacteriophages, p. 475-520. In R. Calendar (ed.), The bacteriophages, vol. 2. Plenum Press, New York. 100. Mitra, A. Personal communication. 101. Moestrup, O., and H. A. Thomsen. 1974. An ultrastructural study of the flagellate Pyramimonas orientalis with particular emphasis on Golgi apparatus activity and the flagellar apparatus. Protoplasma 81:247-269. 102. Moss, B. 1990. Regulation of vaccinia virus transcription. Annu. Rev. Biochem. 59:661-688. 103. Muller, D. G., H. Kawai, B. Stacke, and S. Lanka. 1990. A virus infection in the marine brown alga Ectocarpus siliculosus (Phaeophyceae). Bot. Acta 103:72-82. 104. Muller, D. G., and S. Lanka. Personal communication. 105. Muscatine, L. 1971. Experiments on green algae coexistent with zooxanthellae in sea anemones. Pac. Sci. 25:13-21. 106. Muscatine, L., C. B. Cook, R. L. Pardy, and R. R. Pool. 1975. Uptake, recognition and maintenance of symbiotic Chlorella
by Hydra viridis. Symp. Soc. Exp. Biol. 29:175-203. 107. Nakamura, K., and R. W. Compans. 1979. Host cell- and virus strain-dependent differences in oligosaccharides of hemagglutinin glycoproteins of influenza A viruses. Virology 95:8-23. 108. Narva, K. E., and J. L. Van Etten. Unpublished results. 109. Narva, K. E., J. L. Van Etten, B. E. Slatko, and J. S. Benner. 1988. The amino acid sequence of the eukaryotic DNA [N6adenine]methyltransferase, M.CviBIII, has regions of similarity with the prokaryotic isoschizomer M.TaqI and other DNA [N6-adenine]methyltransferases. Gene 74:253-259. 110. Narva, K. E., D. L. Wendell, M. P. Skrdla, and J. L. Van Etten. 1987. Molecular cloning and characterization of the gene encoding the DNA methyltransferase, M.CviBIII, from Chlorella virus NC-1A. Nucleic Acids Res. 15:9807-9823. 111. Nichols, H. W., and H. C. Bold. 1965. Trichosarcina polymorpha gen. et. sp. nov. J. Phycol. 1:34-38. 112. Nietfeldt, J., K. Lee, and J. L. Van Etten. Chlorella virus PBCV-1 replication is not affected by cytoskeletal disruptors. Intervirology, in press. 113. Nietfeldt, J., and J. L. Van Etten. Unpublished results. 114. Northcote, D. H., K. J. Goulding, and R. W. Horne. 1958. The chemical composition and structure of the cell wall of Chlorella pyrenoidosa. Biochem. J. 70:391-397. 115. O'Brien, T. L., R. MacLeod, and M. C. MacLean. 1984. Absence of lytic virus in two species of symbiotic algae within the sea anemone Anthopleura xanthogrammica (Coelenterata: Anthozoa). Trans. Am. Microsc. Soc. 103:228-232. 116. Oh-Hama, T., and S. Miyachi. 1988. Chlorella, p. 3-26. In M. A. Borowitzka and L. J. Borowitzka (ed.), Micro-algal biotechnology. Cambridge University Press, Cambridge. 117. Oliviera, L., and T. Bisalputra. 1978. A virus infection in the brown alga Sorocarpus uvaeformis (Lyngbye) Pringsheim (Phaeophyta, Ectocarpales). Ann. Bot. 42:439-445. 118. Ortin, J., L. Enjuanes, and E. Vinuela. 1979. Cross-links in African swine fever virus DNA. J. Virol. 31:579-583. 119. Pardy, R. L. Personal communication. 120. Pardy, R. L. 1974. Some factors affecting the growth and distribution of the algal endosymbionts of Hydra viridis. Biol. Bull. 147:105-118. 121. Pardy, R. L. 1983. Preparing aposymbiotic hydra, p. 393-397. In H. M. Lenhoff (ed.), Hydra: research methods. Plenum Press, New York. 122. Pearson, B. R., and R. E. Norris. 1974. Intranuclear virus-like particles in the marine alga Platymonas sp. (Chlorophyta, Prasinophyceae). Phycologia 13:5-9. 123. Pickett-Heaps, J. D. 1972. A possible virus infection in the green alga Oedogonium. J. Phycol. 8:44 47. 124. Pienaar, R. N. 1976. Virus-like particles in three species of phytoplankton from San Juan Island, Washington. Phycologia
15:185-190. 125. Pool, R. R. 1979. The role of algal antigenic determinants in the recognition of potential algal symbionts by cells of chlorohydra. J. Cell Sci. 35:367-379. 126. Posfai, J., A. S. Bhagwat, G. Posfai, and R. J. Roberts. 1989. Predictive motifs derived from cytosine methyltransferases. Nucleic Acids Res. 17:2421-2435. 127. Preisig, H. R., and D. J. Hibberd. 1984. Virus-like particles and endophytic bacteria in Paraphysomonas and Chromophysomonas (Chrysophyceae). Nord. J. Bot. 4:279-285. 128. Que, Q., and J. L. Van Etten. Unpublished results. 129. Reisser, W. 1981. The endosymbiotic unit of Stentor polymorphus and Chlorella sp. Morphological and physiological studies. Protoplasma 105:273-284. 130. Reisser, W. 1986. Endosymbiotic associations of freshwater protozoa and algae. Progr. Protistol. 1:195-214. 131. Reisser, W. 1987. Naturally occurring and artificially established associations of ciliates and algae. Ann. N.Y. Acad. Sci.
503:316-329. 132. Reisser, W., B. Becker, and T. Klein. 1986. Studies on ultrastructure and host range of a Chlorella attacking virus. Protoplasma 135:162-165. 133. Reisser, W., D. E. Burbank, R. H. Meints, B. Becker, and J. L. Van Etten. 1991. Viruses distinguish symbiotic Chlorella spp.
VOL. 55, 1991
of Paramecium bursaria. Endocytobiosis Cell Res. 7:245-257. 134. Reisser, W., D. E. Burbank, S. M. Meints, R. H. Meints, B. Becker, and J. L. Van Etten. 1988. A comparison of viruses infecting two different Chlorella-like green algae. Virology 167:143-149. 135. Reisser, W., D. Fischer-Defoy, J. Staudinger, N. Schilling, and K. Hausmann. 1984. The endosymbiotic unit of Climacostomum virens and Chlorella sp. I. Morphological and physiological studies on the algal partner and its localization in the host cell. Protoplasma 119:93-99. 136. Reisser, W., T. Klein, and B. Becker. 1988. Studies on phycoviruses. I. On the ecology of viruses attacking Chlorellae exsymbiotic from an European strain of Paramecium bursaria. Arch. Hydrobiol. 111:575-583. 137. Richmond, A. (ed.). 1986. Handbook of microalgal mass culture. CRC Press, Inc., Boca Raton, Fla. 138. Rohozinski, J., L. E. Girton, and J. L. Van Etten. 1989. Chlorella viruses contain linear nonpermuted double stranded DNA genomes with covalently closed hairpin ends. Virology 168:363-369. 139. Rohozinski, J., and J. L. Van Etten. 1989. Characterization of DNA polymerases in an uninfected and virus PBCV-1 infected green alga-Chlorella strain NC64A. Intervirology 30:156-162. 140. Rohozinski, J., and J. L. Van Etten. 1989. Purification and characterization of soluble and membrane-associated DNA polymerases from a virus PBCV-1 infected green alga Chlorella NC64A. J. Exp. Bot. 40:1293-1298. 141. Rosowski, J. R., and B. C. Parker (ed.). 1982. Selected papers in phycology II. Phycological Society of America, Inc., Lawrence, Kans. 142. Schlesinger, M. J., and S. Schlesinger. 1987. Domains of virus glycoproteins. Adv. Virus. Res. 33:1-44. 143. Schnepf, E., C. J. Soeder, and E. Hegewald. 1970. Polyhedral virus like particles lysing the aquatic Phycomycete Aphelidium sp., a parasite of the green alga Scenedesmus armatus. Virology 42:482-487. 144. Schuster, A. M., D. E. Burbank, B. Meister, M. P. Skrdla, R. H. Meints, S. Hattman, D. Swinton, and J. L. Van Etten. 1986. Characterization of viruses infecting a eukaryotic Chlorellalike green alga. Virology 150:170-177. 145. Schuster, A. M., L. Girton, D. E. Burbank, and J. L. Van Etten. 1986. Infection of a Chlorella-like alga with the virus PBCV-1: transcriptional studies. Virology 148:181-189. 146. Schuster, A. M., M. Graves, K. Korth, M. Ziegelbein, J. Brumbaugh, D. Grone, and R. H. Meints. 1990. Transcription and sequence studies of a 4.3-kbp fragment from a dsDNA eukaryotic algal virus. Virology 176:515-523. 147. Schuster, A. M., J. A. Waddle, K. Korth, and R. H. Meints. 1990. The chloroplast genome of an exsymbiotic Chlorella-like green alga. Plant Mol. Biol. 14:859-862. 148. Shelef, G., and C. J. Soeder (ed.). 1980. Algal biomass: production and use. Elsevier/North-Holland Biomedical Press, Amsterdam. 149. Sherman, L. A., and R. M. Brown. 1978. Cyanophages and viruses of eukaryotic algae, p. 145-234. In H. Fraenkel-Conrat and R. R. Wagner (ed.), Comprehensive Virology, vol. 12. Plenum Press, New York. 150. Shields, S. L., D. E. Burbank, R. Grabherr, and J. L. Van Etten. 1990. Cloning and sequencing the cytosine methyltransferase gene M.CviJI from Chlorella virus IL-3A. Virology 176:16-24. 151. Shihra, I., and R. W. Krauss. 1965. Chlorella: physiology and taxonomy of forty one isolates. University of Maryland Press, College Park, Md. 152. Sicko-Goad, L., and G. Walker. 1979. Viroplasm and large virus-like particles in the dinoflagellate Gymnodinium uberrimum. Protoplasma 99:203-210. 153. Sieburth, J. M., P. W. Johnson, and P. E. Hargraves. 1988. Ultrastructure and ecology of Aureococcus anophagefferens gen. et sp. nov. (Chrysophyceae): the dominant picoplankter during a bloom in Narragansett bay, Rhode Island, summer 1985. J. Phycol. 24:416-425. 154. Siegel, R. W. 1960. Hereditary endosymbiosis in Paramecium
VIRUSES OF EUKARYOTIC ALGAE
619
bursaria. Exp. Cell Res. 19:239-252. 155. Skotnicki, A., A. Gibbs, and N. G. Wrigley. 1976. Further studies on Chara corrallina virus. Virology 75:457-468. 156. Skrdla, M. P., D. E. Burbank, Y. Xia, R. H. Meints, and J. L. Van Etten. 1984. Structural proteins and lipids in a virus, PBCV-1, which replicates in a Chlorella-like alga. Virology 135:308-315. 157. Skrdla, M. P., M. Graves, J. L. Van Etten, and R. H. Meints. Unpublished results. 158. Slatko, B. E., J. S. Benner, T. Jager-Quinton, L. S. Moran, T. G. Simcox, E. M. Van Cott, and G. G. Wilson. 1987. Cloning, sequencing and expression of the TaqI restrictionmodification system. Nucleic Acids Res. 15:9781-9796. 159. Smith, D. C., and A. E. Douglas. 1987. The biology of symbiosis. Edward Arnold, Ltd., London. 160. Smith, H. O., and S. V. Kelly. 1984. Methylases of the type II restriction-modification systems, p. 39-71. In A. Razin, H. Cedar, and A. D. Riggs (ed.), DNA methylation: biochemistry and biological significance. Springer-Verlag KG, Berlin. 161. Sogo, J. M., J. M. Almendral, A. Talavera, and E. Vinuela. 1984. Terminal and internal inverted repetitions in African swine fever virus DNA. Virology 133:271-275. 162. Soyer, M. 0. 1978. Particules de type viral et filaments trichocystoides chez les Dinoflagelles. Protistologica 14:53-58. 163. Stanker, L. H., and L. R. Hoffman. 1979. A simple histological assay to detect virus-like particles in the green alga Cylindrocapsa (Chlorophyta). Can. J. Bot. 57:838-842. 164. Stanker, L. H., L. R. Hoffman, and R. MacLeod. 1981. Isolation and partial chemical characterization of a virus-like particle from a eukaryotic alga. Virology 114:357-369. 165. Stefan, C., Y. Xia, and J. L. Van Etten. 1991. Molecular cloning and characterization of the gene encoding the adenine methyltransferase M.CviRI from Chlorella virus XZ-6E. Nucleic Acids Res. 19:307-311. 166. Stelker, S., Y. Xia, and J. L. Van Etten. Unpublished results. 167. Strasser, P., Y. Zhang, J. Rohozinski, and J. L. Van Etten. 1991. The termini of the Chlorella virus PBCV-1 genome are identical 2.2-kbp inverted repeats. Virology 180:763-769. 168. Suttle, C. A. Personal communication. 169. Suttle, C. A., A. M. Chan, and M. T. Cottrell. 1990. Infection of phytoplankton by viruses and reduction of primary productivity. Nature (London) 347:467-469. 170. Swale, E. M. F., and J. H. Belcher. 1973. A light and electron microscope study of the colourless flagellate Aulacomonas Skuja. Arch. Mikrobiol. 92:91-103. 171. Takeda, H. 1988. Classification of Chlorella strains by cell wall sugar composition. Phytochemistry 27:3822-3826. 172. Takeda, H. 1988. Classification of Chlorella strains by means of the sugar components of the cell wall. Biochem. Syst. Ecol. 16:367-371. 173. Tessman, I. 1985. Genetic recombination of the DNA plant virus PBCV-1 in a Chlorella-like alga. Virology 145:319-322. 174. Tikhonenko, A. S., and N. B. Zavarzina. 1966. Morphology of the lytic agents of Chlorella pyrenoidosa Mikrobiologiya 35: 850-852. 175. Tomas, R. N., and E. R. Cox. 1973. Observations on the symbiosis of Peridinium balticum and its intracellular alga. I. Ultrastructure. J. Phycol. 9:304-323. 176. Toth, R., and R. T. Wilce. 1972. Viruslike particles in the marine alga Chorda tomentosa Lyngbye (Phaeophyceae). J. Phycol. 8:126-130. 177. Traylor, E., Y. Xia, P. Roy, and J. L. Van Etten. Unpublished results. 178. Trench, R. K. 1979. The cell biology of plant-animal symbiosis. Annu. Rev. Plant Physiol. 30:485-531. 179. Van Etten, J. L. Unpublished results. 180. Van Etten, J. L., D. E. Burbank, J. Joshi, and R. H. Meints. 1984. DNA synthesis in a Chlorella-like alga following infection with the virus PBCV-1. Virology 134:443-449. 181. Van Etten, J. L., D. E. Burbank, D. Kuczmarski, and R. H. Meints. 1983. Virus infection of culturable Chlorella-like algae and development of a plaque assay. Science 219:994-996. 182. Van Etten, J. L., D. E. Burbank, and R. H. Meints. 1986.
620
183. 184. 185. 186.
187.
188. 189.
190.
191.
192.
193. 194. 195. 196.
197. 198. 199.
200. 201. 202.
203. 204.
205.
VAN ETTEN ET AL.
Replication of the algal virus PBCV-1 in UV-irradiated Chlorella. Intervirology 26:115-120. Van Etten, J. L., D. E. Burbank, A. M. Schuster, and R. H. Meints. 1985. Lytic viruses infecting a Chlorella-like alga. Virology 140:135-143. Van Etten, J. L., D. E. Burbank, Y. Xia, and R. H. Meints. 1983. Growth cycle of a virus, PBCV-1, that infects Chlorellalike algae. Virology 126:117-125. Van Etten, J. L., and R. H. Meints. Unpublished results. Van Etten, J. L., R. H. Meints, D. E. Burbank, D. Kuczmarski, D. A. Cuppels, and L. C. Lane. 1981. Isolation and characterization of a virus from the intracellular green alga symbiotic with Hydra viridis. Virology 113:704-711. Van Etten, J. L., R. H. Meints, D. Kuczmarski, D. E. Burbank, and K. Lee. 1982. Viruses of symbiotic Chlorella-like algae isolated from Paramecium bursaria and Hydra viridis. Proc. Natl. Acad. Sci. USA 79:3867-3871. Van Etten, J. L., Q. Que, L. C. Lane, and M. K. Brakke. Unpublished results. Van Etten, J. L., A. M. Schuster, L. Girton, D. E. Burbank, D. Swinton, and S. Hattman. 1985. DNA methylation of viruses infecting a eukaryotic Chlorella-like green alga. Nucleic Acids Res. 13:3471-3478. Van Etten, J. L., A. M. Schuster, and R. H. Meints. 1988. Viruses of eukaryotic Chlorella-like algae, p. 411-428. In Y. Koltin and M. J. Leibowitz (ed.), Viruses of fungi and simple eukaryotes. Marcel Dekker, Inc., New York. Van Etten, J. L., C. H. Van Etten, J. K. Johnson, and D. E. Burbank. 1985. A survey for viruses from freshwater that infect a eukaryotic Chlorella-like green alga. Appl. Environ. Microbiol. 49:1326-1328. Van Etten, J. L., Y. Xia, and R. H. Meints. 1987. Viruses of a Chlorella-like green alga, p. 307-325. In T. Kosuge and E. W. Nester (ed.), Plant-microbe interactions, vol. II. Macmillan Publishing Co., New York. Vivier, E., A. Petitprez, and A. F. Chive. 1967. Observations ultrastructurales sur les Chlorelles symbiotes de Paramecium bursaria. Protistologica 3:325-334. Waddle, J. A., R. H. Meints, D. E. Burbank, and J. L. Van Etten. Unpublished results. Waddle, J. A., A. M. Schuster, K. W. Lee, and R. H. Meints. 1990. The mitochondrial genome of an exsymbiotic Chlorellalike green alga. Plant Mol. Biol. 14:187-195. Wang, I., M. Graves, Y. Li, L. C. Lane, R. H. Meints, and J. L. Van Etten. Unpublished results. Wang, I., L. C. Lane, Y. Li, and J. L. Van Etten. Unpublished results. Wang, I., Q. Que, and J. L. Van Etten. Unpublished results. Waters, R. E., and A. T. Chan. 1982. Micromonas pusilla virus: the virus growth cycle and associated physiological events within the host cells; host range mutation. J. Gen. Virol. 63:199-206. Weis, D. S. 1977. Synchronous development of symbiotic Chlorellae within Paramecium bursaria. Trans. Am. Microsc. Soc. 96:82-86. Weis, D. S. 1978. Correlation of infectivity and concanavalin A agglutinability of algae exsymbiotic from Paramecium bursaria. J. Protozool. 25:366-370. Wiggins, B. A., and M. Alexander. 1985. Minimum bacterial density for bacteriophage replication: implications for significance of bacteriophages in natural ecosystems. Appl. Environ. Microbiol. 49:19-23. Willis, D. B. (ed.). 1985. Iridoviridae. Curr. Top. Microbiol. Immunol. 116:1-173. Willis, D. B., R. Goorha, and A. Granoff. 1984. DNA methyltransferase induced by frog virus 3. J. Virol. 49:86-91. Willis, D. B., and A. Granoff. 1980. Frog virus 3 is heavily
MICROBIOL. REV.
methylated at CpG sequences. Virology 107:250-257. 206. Wilson, G. C. 1988. Type II restriction-modification systems. Trends Genet. 4:314-318. 207. Wittek, R., A. Menna, H. K. Muller, D. Schumperli, P. G. Boseley, and R. Wyler. 1978. Inverted terminal repeats in rabbit poxvirus and vaccinia virus DNA. J. Virol. 28:171-181. 208. Wittek, R., and B. Moss. 1980. Tandem repeats within the inverted terminal repetition of vaccinia virus DNA. Cell 21: 277-284. 209. Xia, Y., D. E. Burbank, L. Uher, D. Rabussay, and J. L. Van Etten. 1986. Restriction endonuclease activity induced by PBCV-1 virus infection of a Chlorella-like green alga. Mol. Cell. Biol. 6:1430-1439. 210. Xia, Y., D. E. Burbank, L. Uher, D. Rabussay, and J. L. Van Etten. 1987. IL-3A virus infection of a Chlorella-like green alga induces a DNA restriction endonuclease with novel sequence specificity. Nucleic Acids Res. 15:6075-6090. 211. Xia, Y., D. E. Burbank, and J. L. Van Etten. 1986. Restriction endonuclease activity induced by NC-1A virus infection of a Chlorella-like green alga. Nucleic Acids Res. 14:6017-6030. 212. Xia, Y., R. Morgan, I. Schildkraut, and J. L. Van Etten. 1988. A site-specific single strand endonuclease activity induced by NYs-1 virus infection of a Chlorella-like green alga. Nucleic Acids Res. 16:9477-9487. 213. Xia, Y., K. E. Narva, and J. L. Van Etten. 1987. The cleavage site of the RsaI isoschizomer, CviII, is G/ITAC. Nucleic Acids Res. 15:10063. 214. Xia, Y., M. P. Skrdla, and J. L. Van Etten. Unpublished
results. 215. Xia, Y., and J. L. Van Etten. Unpublished results. 216. Xia, Y., and J. L. Van Etten. 1986. DNA methyltransferase induced by PBCV-1 virus infection of a Chlorella-like green alga. Mol. Cell. Biol. 6:1440-1445. 217. Yamada, T. 1982. Isolation and characterization of chloroplast DNA from Chlorella ellipsoidea. Plant Physiol. 70:92-96. 218. Yamada, T. 1983. Characterization of inverted repeat sequences and ribosomal RNA genes of chloroplast DNA from Chlorella ellipsoidea. Curr. Genet. 7:481-487. 219. Yamada, T., and K. Sakaguchi. 1982. Comparative studies on Chlorella cell walls: induction of protoplast formation. Arch. Microbiol. 132:10-13. 220. Yamada, T., and K. Sakaguchi. 1982. Electron microscopic studies of Chlorella ellipsoidea protoplast formation. J. Gen. Microbiol. 128:1319-1327. 221. Yao, M. C., and C. H. Yao. 1981. Repeated hexanucleotide C-C-C-C-A-A is present near free ends of macronuclear DNA of Tetrahymena. Proc. Natl. Acad. Sci. USA 78:7436-7439. 222. Yonker, C. R., K. D. Caldwell, J. C. Giddings, and J. L. Van Etten. 1985. Physical characterization of PBCV virus by sedimentation field flow fractionation. J. Virol. Methods 11:145160. 223. Yuen, L., and B. Moss. 1987. Oligonucleotide sequence signaling transcriptional termination of vaccinia virus early genes. Proc. Natl. Acad. Sci. USA 84:6417-6421. 224. Zavarzina, N. B. 1961. A lytic agent in cultures of Chlorella pyrenoidosa Pringh. Dokl. Akad. Nauk SSSR Ser. Biol. 137: 435-437. 225. Zavarzina, N. B. 1964. Lysis of Chlorella cultures in the absence of bacteria. Mikrobiologiya 33:561-564. 226. Zavarzina, N. B., and A. E. Protsenko. 1958. The lysis of Chlorella pyrenoidosa Pringh cultures. Dokl. Akad. Nauk SSSR Ser. Biol. 122:936-939. 227. Zhang, Y., D. E. Burbank, and J. L. Van Etten. 1988. Chlorella viruses isolated in China. Appl. Environ. Microbiol. 54:21702173. 228. Zhang, Y., and J. L. Van Etten. Unpublished results.
MICROBIOLOGICAL REVIEWS, Dec. 1991, p. 586-620 0146-0749/91/040586-35$02.00/0 Copyright C 1991, American Society for Microbiology
Viruses and Viruslike Particles of Eukaryotic Algae JAMES L. VAN ETTEN,1* LESLIE C. LANE,' AND RUSSEL H. MEINTS2 Department of Plant Pathology, University of Nebraska, Lincoln, Nebraska 68583-0722,1 and Department of Botany and Plant Pathology, Center for Gene Research and Biotechnology, Oregon State University, Corvallis, Oregon 97331-2906 INTRODUCTION ...................................................................................
586
CHARACTERISTICS OF CHLORELLA SPECIES
587
......................................................................
Properties of Chlorella Species ...................................................................................587 590 Symbiotic Chlorella Species ................................................................................... Chlorella Strains NC64A, Nla, Pbi, and SAG-241-80 .................................................................591 General
DISCOVERY OF CHLORELLA VIRUSES ................................................................................. 591 Initial Observations
Development of
a
591
...................................................................................
Culturable
System
for
PBCV-1
...................................................................... 592
PROPERTIES OF CHLORELLA VIRUSES ................................................................................ 592 General Characteristics of
Composition
PBCV-1
of
PBCV-1
PBCV-1
594
Genome ...................................................................................
PBCV-1 Chlorella Strain NC64A Viruses Viruses Infecting Chlorella Strain Pbi REPLICATION OF PBCV-1 Attachment of PBCV-1 to Chlorella Strain NC64A Intracellular Site of PBCV-1 Replication Effect of PBCV-1 Infection Host DNA, RNA, and TRANSCRIPTION OF PBCV-1 Serology
592
...................................................................................
...................................................................................
of
596
597
...................................................................................
Additional
..............................................................................
598 598
...................................................................................
598
................................................................................... ....................................................................
598
.................................................................................
600
on
Protein
Syntheses .........................................604 604
...................................................................................
VIRUSES ENCODE DNA METHYLTRANSFERASES AND DNA SITE-SPECIFIC (RESTRICTION) ENDONUCLEASES
PBCV-1
607
...................................................................................
Methyltransferases and DNA Site-Specific Endonucleases ........................................607 Viral DNA Methyltransferases and DNA Site-Specffic Endonucleases ..............................607
DNA
Additional
SEQUENCE OF VIRUS-ENCODED DNA METHYLTRANSFERASES ............................................609
FUNCTION OF VIRUS-ENCODED DNA SITE-SPECIFIC ENDONUCLEASES AND DNA METHYLTRANSFERASES ...................................................................................
609
NATURAL HISTORY OF THE CHLORELLA VIRUSES ..............................................................610 COMPARISON OF CHLORELLA VIRUSES WITH OTHER VIRUSES ...........................................611
PROPERTIES OF VIRUSES OR VLPs OF OTHER ALGAE
615
ACKNOWLEDGMENTS ...................................................................................
616
REFERENCES ...................................................................................
616
photosynthesis, cell cycle regulation, cell motility, and nonMendelian (chloroplast) inheritance. Given the importance of eukaryotic algae, there is surprisingly little information about viruses or viruslike particles (VLP) in these organ-
INTRODUCTION
Eukaryotic algae range in size and complexity from small (microalgae) to large multicellular organisms (macroalgae), such as seaweeds. Most eukaryotic algae are free living, and members can grow and survive in many terrestrial and aquatic environments. A few, however, are found only in symbiotic association with other organisms such as molluscs, flatworms, coelenterates, protozoa, and fungi (lichens). Algae significantly influence aquatic environments, both as primary producers in the food chain and, conversely, as pollutants when growth becomes uncontrolled. Because most algae require only light, simple salts, and a nitrogen source for rapid growth, they are potential producers of proteins, carbohydrates, and fats for feeding humans and animals. In addition, algae are a source of useful compounds such as 13-carotene, alginic acid, carrageenans, and agar (12, 137, 148). Algae such as Chlamydomonas and Chlorella spp. have also served as models for studying unicellular organisms
*
612
........................................................
CONCLUDING COMMENTS ...................................................................................
isms. It is difficult to credit the first person who described a virus or VLP in a eukaryotic alga. The problem is complicated because early reports consisted solely of microscopic observations and, in some cases, cultures were not axenic. A few papers in the Russian literature as long as 30 years ago described a lytic activity in cultures of the green alga Chlorella pyrenoidosa (174, 224-226). This lytic activity was given the name "chlorellophage." However, these investigators acknowledged that their cultures were contaminated with bacteria. Furthermore, the chlorellophage had typical bacteriophage morphology (174, 224). Therefore, despite the name chlorellophage, the host for these particles is unclear. Several investigators, almost simultaneously, reported the presence of VLPs in eukaryotic algae in the early 1970s. Lee (73) described 50- to 60-nm polyhedral particles in vegetative cells of the red alga Sirodotia tenuissima, Pickett-Heaps
Corresponding author. 586
VOL. 55, 1991
(123) described polyhedral particles (240 nm in diameter) in germlings of the green alga Oedogonium sp., and Toth and Wilce (176) described 170-nm polyhedral particles in spores of the marine brown alga Chorda tomentosa. Since these initial reports, VLPs or viruses have been reported in at least 44 taxa of eukaryotic algae (Table 1). These are distributed in 10 of the 14 recognized classes of eukaryotic algae (141). Particles have not been found in any member of the families Euglenophyceae, Bacillariophyceae, Raphidophyceae, and Tribophyceae. VLPs and viruses of eukaryotic algae have been reviewed previously by Lemke (74), Sherman and Brown (149), Dodds (27, 28), Martin and Benson (84), and Van Etten et al. (190, 192). The review by Sherman and Brown contains several electron micrographs of these particles, including some not published elsewhere. One other report is noteworthy. In 1970 Schnepf et al. (143) described large (200 nm in diameter) polyhedral particles in an Aphelidium sp. (which, although described as a fungus, may actually be a protozoan) that parasitized the green alga Scenedesmus armatus. These particles were much larger than most fungal viruses (68) but were similar in size and morphology to VLPs subsequently found in eukaryotic algae. With few exceptions, the reports of VLPs in eukaryotic algae typically consist of single accounts of microscopic observations. Furthermore, many of them were with fieldcollected algae, which are not available for further study. Characterization of these particles was not pursued because of low concentrations. Several factors contributed to the low concentrations: (i) usually only a few algal cells contained particles, (ii) usually the cells contained particles only at one stage of the algal life cycle, (iii) usually cells containing particles did not lyse, and (iv) in most cases the particles were not infectious. In addition, some of the algae could not be cultured and others were inconvenient to work with (e.g., filamentous algae). However, these difficulties are no longer a problem for a family of viruses which infect and replicate in certain strains of unicellular Chlorella-like green algae. The first of the Chlorella viruses was discovered about 10 years ago in Chlorella-like algae (zoochlorella) symbiotic with Hydra viridis (96, 186, 187) and subsequently in Chlorella-like algae in Paramecium bursaria (132, 181, 183, 187). The algae from P. bursaria can be grown free of the paramecium, and the cultured Chlorella strains NC64A and Pbi or their equivalents serve as hosts for many viruses. These viruses can be produced in large quantities, assayed by plaque formation, and analyzed by standard bacteriophage techniques. This review will describe the biology of the Chlorella virus hosts, which normally exist as symbionts, and then focus on the discovery, biology, and molecular biology of the Chlorella viruses. Partially characterized viruses or VLPs of other eukaryotic algae are discussed at the end of the review. CHARACTERISTICS OF CHLORELLA SPECIES General Properties of Chlorella Species The genus Chlorella contains small spherical or ellipsoidal, unicellular, nonmotile, asexually reproducing green algae. Chlorella species have a rigid cell wall and typically contain a single chloroplast, which may or may not contain a pyrenoid. They have a simple developmental cycle and reproduce by mitotic division. Vegetative cells increase in size and divide into two, four, eight, or more progeny (called autospores), which are released by rupture or enzymatic
VIRUSES OF EUKARYOTIC ALGAE
587
digestion of the parental walls. The autospores develop rapidly into vegetative cells. The number of progeny formed from a single vegetative cell varies with species and environmental conditions (200). Chlorella species typically grow on a simple salts solution and utilize nitrate, ammonium, or amino nitrogen as a nitrogen source (116). However, they often grow best on casein hydrolysate (91, 151). Simple sugars such as sucrose or glucose increase the growth rate of some species but inhibit the growth of others. The doubling time varies with the strain or species, but many of them double in about 24 h. Algae assigned to the genus Chlorella are more heterogeneous than their simple morphology suggests (25, 58, 65, 151). Huss et al. are examining the relationships among Chlorella species by using DNA base ratios, DNA hybridization, and DNA reassociation kinetics (see reference 58 and references cited therein). The guanosine-plus-cytosine content of nuclear DNA, which ranges from 43 to 79% (49), reflects the heterogeneity of algae assigned to the genus Chlorella. However, most isolates assigned to the same species have similar G+C contents. The Chlorella base compositions were determined either by measuring DNA melting temperatures or by performing CsCl density equilibrium centrifugation. Recently, high levels of 5-methylcytosine (5mC) were found in the nuclear DNAs of Chlorella strain NC64A (21% of the cytosines were methylated [189]) and Chlorella sorokiniana (16% of the cytosines were methylated [22]). If high concentrations of 5mC are a common feature of Chlorella DNAs, many of the published Chlorella G+C contents may be low because 5mC can lower DNA melting temperatures and buoyant densities. The DNA content of Chlorella cells was reported to range from about 15 fg per cell for cultured Chlorella strains 211/le and NC64A to 47 fg per cell for symbiotic Chlorella strains isolated from hydras (88). We estimate that Chlorella strain NC64A contains about 40 fg (40 Mbp) of DNA per cell (180), which is almost three times higher than that found by McAuley and Muscatine (88). Assuming the higher value and assuming haploidy, the Chlorella strain NC64A genome is about twice the size of the Saccharomyces cerevisiae genome and one-half the size of the Arabidopsis thaliana genome. Repetitive sequences make up only a small fraction (about 5 to 8%) of the total Chlorella DNA (58). The cell wall composition of Chlorella species also varies widely (see, e.g., references 3, 76, 114, 171, 172, and 219). Typically, Chlorella cell walls contain polysaccharides with a variety of component sugars as well as lesser amounts of protein and lipid. In addition, walls of some Chlorella species contain a layer of sporopollenin (3), a polymer of carotenoid derivatives resistant to most chemicals (13). The difficulty of classifying Chlorella species is illustrated by a report that the chemical compositions of the cell walls of some isolates assigned to the same species differ markedly, e.g., for Chlorella vulgaris and Chlorella ellipsoidea (219). Because of the heterogeneity in the genus Chlorella, properties discovered in one Chlorella species or strain may not always apply to another. As noted below, viruses can distinguish at least two exsymbiotic Chlorella strains from each other and from other free-living Chlorella species. If viruses infecting additional Chlorella species are found, viruses could be useful in classifying Chlorella species in the same way that bacteriophages are useful in bacterial classification (see, e.g., reference 2).
588
VAN ETTEN ET AL.
MICROBIOL. REV. TABLE 1. Viruses and VLPs of eukaryotic algae
Host species'
Location of virogenic stroma
Infection and replication facts
Morphological
Nucleic acid
characteristics
Key references
Rhodophyceae
Porphyridium purpureumb
Unkc
Polyhedral, 40 nm in diameter
Unk
18
Unk
Polyhedral, 50-60 nm in diameter
Unk
73
Nucleus and cytoplasm
Unk
Two types of particles were observed in the same cell: (i) polyhedral, 99 nm in diameter; (ii) stalked particle, head 120 nm diameter, length 240 nm; surrounded by a membrane
Unk
124
Dinophyceae Gymnodinium uberrimunf *Gyrodinium resplendensg
Cytoplasm Cytoplasm
Unk Unk
Polyhedral, 385 nm in diameter Two types of particles: (i) tubular arrays, with particles 35 nm in diameter; (ii) particles 20 nm in diameter
Unk Unk
152 34, 162
Chrysophyceae Chromophysomonas cor-
Cytoplasm
Unk
Unk
127
Dinobryon sp.J
Cytoplasm Cytoplasm
Unk Unk
Two polyhedral particles, 50-60 nm and 150-180 nm, in diameter Polyhedral, 35 nm in diameter Polyhedral, 100 nm in diameter
Unk Unk
Hydrurus foetidusk Mallomonas sp. Paraphysomonas caelifri-
Cytoplasm Cytoplasm Cytoplasm
Unk Unk Unk
Polyhedral, 50-60 nm in diameter Polyhedral, 175 nm in diameter Polyhedral, 150-180 nm in diame-
Unk Unk Unk
175 Carson and Brown (unpublished) cited in 149 55 152 127
Paraphysomonas corynephora'
Cytoplasm
Unk
Unk
127
Paraphysomonas bourrellyi
Cytoplasm
Unk
Unk
127
Unk Unk
Unk Unk
Polyhedral, 22 nm in diameter Polyhedral, 22 nm in diameter
Unk Unk
82 82
Nucleus?
Unk Unk
Polyhedral, 65 nm in diameter Polyhedral, 400 nm in diameter
Unk Unk
Sirodotia tenuissimad Cryptophyceae *Cryptomonas sp.'
Nucleus and cytoplasm Cytoplasm
nutah
Chrysophycean-like'
ca'
Prymnesiophyceae (Haptophyceae) *Chrysochromulina sp. *Coccolithus huxleyi *Hymenomonas carterae Strain ln
Cytoplasm
Strain 2
ter Three polyhedral particles, 50-60 nm, 150-180 nm, and 270-300 nm in diameter Two polyhedral particles, 50-60 nm and 150-180 nm in diameter
124
Cooper and Brown (unpublished) cited in 149
Eustigmatophyceae Monodus sp.°
Nucleus?
Unk
Polyhedral, 400 nm in diameter
Unk
Huang and Hommersand (unpublished) cited in 149
Phaeophyceae *Chorda tomentosaP *Ectocarpus fasciculatisq
Cytoplasm? Nucleus?
Unk Unk
Polyhedral, 170 Polyhedral, 170
nm nm
in diameter in diameter
Unk Leads to cell lysis Infects swimming zoo-
176 4, 21
*Ectocarpus
siliculosusr
Cytoplasm
dsDNA
Polyhedral, 130
nm
in diameter
*Feldmannia
sp.S
Nucleus?
dsDNA
Polyhedral, 150
nm
in diameter
spores
Replicates only in
*Pylaiella littoralis'
*Sorocarpus uvaeformisu *Streblonema sp.v
103 51
meiotic
sporangia
Nucleus then cytoplasm?
Unk
Cytoplasm Cytoplasm
Unk Unk
Polyhedral, 130-170
nm
in diame-
Unk
83
Infectious Unk
117 71
ter
Polyhedral, 170 nm in diameter Polyhedral, 135-150 nm in diameter
Continued on following page
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
589
TABLE 1-Continued Host specieSa
Prasinophyceae *Heteromastix sp.
Location of virogenic stroma
Nucleus?
Nucleic acid
Unk
Morphological
characteristics
Polyhedral, 320 nm, with complex membranes
Infection and replication facts
Unk
KeY references e
Ott and Hommersand (un-
published) Mesostigma viridew *Micromonas pusillax
Cytoplasm
*Platymonas sp.Y *Pyramimonas orientalisz
Cytoplasm
Unk dsDNA
Polyhedral, 130 nm in diameter Polyhedral, 130-135 nm in diameter
Nucleus Nucleus
Unk Unk
Polyhedral, 51-58 nm in diameter Two polyhedral particles, 60 and 200 nm in diameter
Chlorophyceae Aulacomonas Sp.aa
Cytoplasm?
Unk
*Brachiomonas sp.
Cytoplasm?
Unk
Chlorella spp.bb
Unk
Unk
Chlorella spp.Cc Chlorella strain NC64Add
Cytoplasm
Cytoplasm
dsDNA dsDNA
Polyhedral, 200-230 nm in diameter, with tail 150-200 nm long Polyhedral, 380-400 nm in diameter; some had long tails (500 nm) Polyhedral head, 41 nm in diameter, with a 24-nm-long, 10-nmwide tail Polyhedral, 170-180 nm Polyhedral, 150-190 nm
Chlorella strain Pbiee
Cytoplasm
dsDNA
Polyhedral, 140-150 nm
Unk Virus adheres to the surface of host cells, not known if uncoating occurs at the surface or internally Unk Unk
cited in 149 97 87, 124, 168, 199
122 101
Unk
170
Unk
54
Unk
174
Unk Attaches specifically to host walls and uncoats at surface Attaches specifically to host walls and un-
96, 186, 187 63, 181, 184, 187
132, 134
coats at sur-
Chlorococcum minutumrf
Cytoplasm
dsDNA
Polyhedral, 180-220 nm in diameter, has a tail
Cylindrocapsa geminellagg
Nucleus, then cytoplasm?
dsDNA
Polyhedral, 200-230 nm in diameter with a membranous coat
Oedogonium sp.h Radiofilum transversale" Stigeoclonium farctum"
Cytoplasm
Polyhedral, 240 nm in diameter Polyhedral, 41 nm in diameter
Chloroplast?
Unk Unk Unk
Uronema gigaskk
Cytoplasm?
dsDNA
Chara corallina"'
Cytoplasm?
ssRNA
Tubular rods, 532 nm long, 18 nm wide, with basic pitch of 2.75 nm
Coleochaete scutata
Nucleus
Unk
Polyhedral, 41 nm in diameter
Nucleus
Tubular? 16.3 nm in cross-section Polyhedral, 390 nm in diameter, some have long tails (1,000 nm)
face Virus infects only zoospores, progeny viruses appear in ca. 8 h, viruses released by lysis of cell Particles may lead to cell lysis Unk Unk Unk Unk
46, 47
56, 163, 164 123 86 86
23, 29, 86
Charophyceae By artificial in38, 155 jection of virus into uninfected Chara cells; chlorosis and death in 10-12 days 86 Unk Continued on following page
590
VAN ETTEN ET AL.
MICROBIOL. REV.
a Classification from reference 141. Asterisks denote marine algae. b The intranuclear particles (termed concentrosomes) are probably not VLPs. The polyhedral particles are present only in the cytoplasm of a small number of cells. c Unk, unknown. d The virogenic stroma is prominent in the apical cell and appears to be transmitted to the daughter cells during division of the apical cell. The host cells with viral particles may be scenescent. f Cells with particles lack a nucleus. g Franca (34) reported two types of VLPs. Soyer (162) suggested that the V2 particles are tricocystoid filaments. The Vl particles do not have helical symmetry and are unlikely to be VLPs. h Two classes of VLPs were sometimes present in the same cell. 'The unidentified chrysophycean alga was an endosymbiont in the dinophycean alga Peridinium balticum. i This VLP may be responsible for rapid cell lysis when natural collections are brought to elevated temperatures of the laboratory, or in late spring when the water temperature increases. This may explain the mass-scale lysis of Dinobryon spp. in major water supplies, giving rise to subsequent blooms of other algae. k VLPs were probably present in zoospores and were not seen in vegetative cells. 'The VLPs may lyse the alga. The VLPs were seen in 75% of the algal cells after enrichment for the alga, which was followed by the disappearance of the alga. m Three size classes of VLPs were observed in the alga; the two smaller particles were occasionally present in the same cell. n Cells containing VLPs were greatly enlarged and contained proliferating chloroplasts. 0 The host could be xanthophycean or intermediate between the Xanthophyceae and Eustigmatophyceae. P VLPs found in settled zoospores of Chorda; spores with VLPs do not synthesize a cell wall and cell organelles were disrupted. e VLPs found in the plurilocular sporangia of native material. VLPs also found in developing zoospores; thus the VLPs may have been transmitted through the zoosporangium to the zoospore by subsequent cleavage. Particles are associated with tubular elements. The particles appear only in gametangia of certain isolates. They have a dsDNA of ca. 350 kbp. No nuclear membrane is present in sporangia with VLPs, but surrounding matrix appears to be nucleoplasm. They have a dsDNA of about 190 kbp. VLPs are present only in sporangia and are not seen in vegetative cells. VLPs are usually present in zoospores but are also seen in vegetative cells of the alga. VLPs are present in vegetative cells. VLPs are present in about 5% of the cells. The alga does not have a cell wall; the particles are infectious. Y Approximately 30% of the cells have particles. Both size classes of VLPs were found in the same cell. aa Aulacomonas sp. is a colorless flagellate, lacking a chloroplast. The VLPs are surrounded by an envelope about 30 nm thick. Smaller polyhedral particles (50 nm in diameter) were present in the nucleus of one cell. bb The presence of viruses in these Chlorella cells is questionable since the cultures were contaminated with bacteria and the particles may be bacteriophage. cc The host Chlorella cell is normally a symbiont of H. viridis. The VLP dsDNA is at least 250 kbp. dd The host Chlorella cell is normally a symbiont of a protozoan, P. bursaria, collected in the United States. The alga can be grown in culture, and a family of viruses which replicate in the alga can be plaque assayed. ee The host Chlorella cell is normally a symbiont of a protozoan, P. bursaria, collected in Europe. The alga can be grown in culture, and a family of viruses which replicate in the alga can be plaque assayed. ff Virus may have a retractable tail. 99 VLPs confined to single-cell germlings, and a heat shock of 40°C, 6-24 h, is necessary to induce particle formation. The heat shock, applied to the zoospores, produces germlings in which 3 to 10% of the cells are infected with VLPs. dsDNA is 275 t8 300 kbp, particles contain at least 10 proteins. hh Detected only in developing germlings. iNo pictures of these particles have been published, but they are reported to resemble the VLPs of Coleochaete spp. S Probably not a VLP. kk About 1% of the cells in the germling or young filament stage contain particles. A heat shock treatment leads to increased VLP prdduction. "The virus was the first experimental infection of a eukaryotic algal cell. Physiochemical data: s2o, = 230S; molecular weight, 3.6 x 106; base ratio of G 24.5, A 28.0, C 20.0, U 27.5; 5% RNA; protein coat molecular weight, ca. 17.5 x 103.
Symbiotic Chlorelia Species Most algae assigned to the genus Chlorella are free living in nature. However, some forms called zoochlorellae or Chlorella-like algae, live as hereditary endosymbionts wvithin freshwater and, to a lesser extent, marine animals (see, e.g., references 40, 105, 159, and 178). For convenience, we will use the term zoochlorellae in this review. Douglas and HIuss (31) have shown that the DNA from several zoochlorellae hybridize with free-living C. vulgaris DNA. Because of the similarity of zoochlorella DNA and C. vulgaris DNA and because some zoochlorellae can be cultured, two groups of investigators (31, 62) have suggested that zoochlorella symbiotic associations may have arisen relatively recently. The green coelenterate H. viridis (106) and the unicellular green protozoan P. bursaria (61) are among the most widely studied organisms containing zoochlorellae. Each gastrodermal cell of H. viridis contains 10 to 20 zoochlorellae and a single hydra contains 1.5 x 10i zoochlorellae under optimum laboratory conditions. P. bursaria typically contains several hundred zoochlorellae. Each algal cell is enclosed within an individual, hostderived vacuole, known as the perialgal vacuole (193). The algae are always located in a well-defined position in the cells of the host hydra or paramecium (159). The zoochlorellae
grow and divide in concert with the host so that the population of zoochlorellae is maintained at a constant level. When green hydras and green paramecia are placed under nongrowing conditions, algal growth also ceases. Thus the zoochlorellae and host grow coordinately (see, e.g., reference 32). Symbiotic zoochlorellae fix CO2 in litht and can transfer 30 to 40% of their total photosynthate to the host hydra or paramecium (33, 98, 131). The symbiotic relationship can also be maintained for at least a short time in dark-grown hydras (120) or paramecia (89). Symbiont-free Hydra viridis can be obtained in the laboratory by growing hydras in glycerol or in high-intensity light either alone or in the presence of a photosynthietic inhibitor such as 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) (121). These treatments produce a white, aposymbiotic hydra. The aposymbiotic hydra can be grown in culture indefinitely if appropriately fed. In contrast, most investigators have had trouble growing hydra zoochlorellae in culture. Although there is one report of the successful culturing of a hydra zoochlorella (59), others believe that this alga may have been a surface contaminant (66).. Aposymbiotic P. bursaria organisms are produced by growth in the dark at high growth rates for several months
VOL. 55, 1991
(130, 154). Paramecium zoochlorellae are apparently easier to culture than those of hydras, since they have been cultured on several occasions (see, e.g., references 75, 119, 154, 201). Cultured zoochlorellae release as much as 80% of their photosynthate to the environment (17), primarily in the form of the disaccharide maltose, whereas free-living Chlorella species do not. Maltose release is pH dependent; it is maximal at about pH 4.0 and declines with increasing pH to nearly zero at pH 7.0. The excreted maltose apparently does not arise directly from starch degradation; instead, Cernichiari et al. (17) suggested that a specific glucosyltransferase condenses glucose at the algal cell surface to produce maltose. The symbiotic relationships of hydras or paramecia and their zoochlorellae can be reestablished by coculture of the host and its symbiont. Symbionts are individually taken up into perialgal vacuoles, which' avoid fusion with lysosomes. The fate of nonsymbiotic Chlorella cells taken in by hydras or paramecia varies with the Chlorella species and host. Typically, the nonsymbiotic Chlorella cells enter food vacuoles and are digested as the vacuoles fuse with lysosomes or else the Chlorella cells fail to grow and divide (see reference 90 and references cited therein). Hereditary endosymbiotic associations are usually, but not always, host specific. For example, reassociation studies with different Chlorella species and aposymbiotic paramecia indicate that the colorless paramecium can take up not only its original symbiotic partners, but also a limited number of other Chlorella species, and can form green paramecia under special conditions. However, only the original paramecium zoochlorellae can reestablish a long-term, stable symbiosis with the protozoan (131). Paramecium zoochlorellae can also form a symbiotic association with hydras which is stable under laboratory conditions (91, 106). Kessler et al. (66) reported that a variety of free-living Chlorella strains could form a symbiotic relationship with hydras. The single characteristic- correlated with this property was the ability of the algae to grow under acidic (pH 4) conditions. This finding, if verified, suggests that specific surface recognition may not be as important in establishing symbiosis as was once thought (see, e.g., references 95 and 125). However, maintaining a stable symbiotic relationship is more complex. The ability of the alga to supply the host with carbon, in the form of maltose (159), and to utilize nitrogen and other host nutrients is probably important. Understanding symbiosis will require discovering the details of the physiological adaptations.
Chlorella Strains NC64A, Nla, Pbi, and SAG-241-80 The hosts for two groups of Chlorella viruses are the exsymbiotic Chlorella strains NC64A and Nla, isolated almost 30 years apart from P. bursaria collected in the United States, and Chlorella strains Pbi and SAG-241-80, isolated from paramecia collected in Europe. Chlorella strains NC64A and Nla have identical mitochondrial and chloroplast genomes (91), and thus the algae are probably identical. We have used NC64A and Nla interchangeably in our virus studies, and to simplify the discussion we will refer to both strains as NC64A in this review. Chlorella strain NC64A can be grown easily in the laboratory on a simple salt solution (Bold's basal medium [111]) modified by adding 0.5% sucrose and 0.1% protease peptone at 25°C in continuous light (ca. 30 microeinsteins m-2 ') with gentle shaking. Under these conditions the alga pro-
VIRUSES OF EUKARYOTIC ALGAE
591
duces four autospores about every 48 h. The algae are oval and measure about 4 by 5 ,um. They have a single cupshaped chloroplast with a prominent pyrenoid. Commercially available enzymes digest the cell walls of some free-living Chlorella species to give viable protoplasts, e.g., C. vulgaris, C. saccharophila, and C. ellipsoidea (1, 7, 220). Chlorella strain NC64A is resistant to commercial cell-wall-degrading enzymes (91). However, we have partially purified an enzyme(s) from virus lysates which degrades Chlorella strain NC64A cell walls. This enzyme(s) (called lysin) can produce protoplasts of Chlorella strain NC64A. As measured by vital stains, the protoplasts live for up to 4 weeks, but attempts to regenerate colonies from these protoplasts have so far been unsuccessful (91). Isolated Chlorella strain NC64A cell walls were analyzed for component monosaccharides (92). Glucose and rhamnose made up 51 and 16% of the total sugars, respectively, with galactose, xylose, arabinose, mannose, and glucosamine each accounting for 5 to 10% of the sugars. These monosaccharides made up about 50% (by weight) of the solubilized wall material. The nature of the remaining wall material is unknown. Chlorella strain NC64A nuclear DNA is 67% G+C (189), and mitochondrial DNA (195) and chloroplast DNA (147) are 32 and 38% G+C, respectively. The nuclear DNA contains methylated bases; SmC makes up 21% of the cytosines, and N6-methyladenine (6mA) makes up 0.6% of the adenines (189). No 4-methylcytosine (4mC) was detected in the nuclear DNA (179). A lambda genomic library of Chlorella strain NC64A (Nla) nuclear DNA has recently been prepared in Escherichia coli K803 (91). The nuclear DNA was not clonable in other E. coli strains, presumably because of its high SmC content (194). A single-copy ,-tubulin gene, which hybridized to a Chlamydomonas tubulin gene, was identified in the Chlorella strain NC64A library and sequenced. The Chlorella tubulin gene has more introns (twelve) than any other known tubulin gene and shares sequence and gene structure with both fungi and higher plants (91). Vectors to transform Chlorella strain NC64A are being constructed with the promoter region of this tubulin gene. Chlorella strain NC64A (Nla) contains a 120-kbp circular chloroplast DNA (147), which is smaller than the 175-kbp chloroplast DNA present in free-living C. ellipsoidea (217, 218). Typically, chloroplast DNAs, including those of C. ellipsoidea, contain an inverted repetitive sequence of about 20 kbp, containing a copy of the 23S, 16S, and 5S rRNA genes. However, Chlorella strain NC64A chloroplast DNA contains only a single rRNA gene. The mitochondrial DNA from Chlorella strain NC64A (Nla) is a 76-kbp circle (195). The mitochondrial DNA was physically mapped, and some of the mitochondrial genes were placed on the map. Beyond these studies, nothing is known about the molecular properties of Chlorella strain NC64A. There is even less information about the molecular properties of Chlorella strains Pbi and SAG-241-80.
DISCOVERY OF CHLORELLA VIRUSES Initial Observations A fortuitous conversation between two of us (R.H.M. and J.V.E.) about 11 years ago led to a simple experiment, which produced an exciting result. We discovered that large (185 nm in diameter) polyhedral VLPs accumulated in zoochlorellae within a few hours after they were isolated from H.
592
VAN ETTEN ET AL.
viridis (Fig. 1). Replication of these particles lysed the entire population of zoochlorellae within 24 h following isolation from hydras (96). These VLPs, which were absent from intracellular zoochlorellae, were named HVCV-1 for H. viridis Chlorella virus. HVCV-1 was isolated and partially characterized. The particles sedimented at 2,600S in sucrose density gradients and had a density of 1.295 g/ml in CsCl. HVCV-1 contained a double-stranded DNA (dsDNA) genome of at least 250 kbp and at least 19 structural proteins varying in apparent size from 10.3 to 82 kDa. The major viral protein of 46 kDa stained with Schiffs reagent, suggesting that it was a glycoprotein (186). Our initial experiments were conducted with zoochlorellae isolated from a hydra strain (Florida strain) which had been maintained in culture for over 20 years. Subsequently we examined zoochlorellae isolated from six other hydra isolates for VLPs. In addition, zoochlorellae were isolated from P. bursaria. In each case, large dsDNA containing VLPs appeared within 24 h of zoochlorella isolation (187). Restriction endonuclease analysis of VLP DNAs revealed four distinct VLPs. The relationship of the VLPs to the source of the hydra was especially interesting. A green hydra obtained from a Massachusetts lake in 1981 yielded a VLP identical to HVCV-1. Green hydras collected in 1981 from lakes in North Carolina, Nebraska, and Massachusetts yielded a second VLP, designated HVCV-2. Two green hydras collected independently in England yielded a third VLP, desig-
nated HVCV-3. The presence of identical genomes in VLPs isolated from zoochlorellae from such diverse geographic locations suggested that the VLPs had a remarkably stable relationship with their zoochlorella host. The source of the VLPs in the hydra-zoochlorellae systems is still unknown. Particles were never observed in intracellular zoochlorellae in the hydras. The particles could result from an external infection which is maintained by the natural death of hydras and release of associated zoochlorellae. Another possibility is that association with hydras suppressed viruses lysogenic with zoochlorellae. These lysogenic viruses might even code for a gene product(s) that mediates symbiosis between the zoochlorellae and the hydras. An additional possibility is that the particles replicate in hydra cells and infect the zoochlorellae during their isolation. Although we never observed VLPs in hydra cells, there is one report of VLPs in a hydra species (Hydra vulgaris) which lacks zoochlorellae (11). These polyhedral particles were smaller (70 to 75 nm) than the VLPs subsequently found in H. viridis zoochlorellae. The possibility of lysogeny in the zoochlorellae or viral replication in the hydras could be determined experimentally; however, these experiments have not been conducted. The DNA restriction fragment pattern of the VLP from the P. bursaria zoochlorellae differed from those of HVCV VLPs, and this VLP was designated PBCV-1 for P. bursaria Chlorella virus. Kawakami and Kawakami (63) had previously observed VLPs infecting zoochlorella in a P. bursaria strain isolated in Japan. Zoochlorellae growing symbiotically inside the paramecia did not contain VLPs. However, the algae became infected immediately after release from the paramecia. The VLPs observed by Kawakami and Kawakami were neither isolated nor physically characterized; however, they were similar in size and morphology to PBCV-1. In summary, VLP infections are common in zoochlorellae shortly after isolation from hydras and paramecia. This can explain the difficulty in culturing zoochlorellae from hydras.
MICROBIOL. REV.
Although the VLP association may contribute to the symbiosis, the relationship in P. bursaria is not as strong, since zoochlorellae have been cultured free of paramecia on several occasions (75, 119, 130, 154, 201). Development of a Culturable System for PBCV-1 We tested several cultured zoochlorellae, reputedly derived from invertebrate hosts, as well as two free-living Chlorella strains, to see whether they could serve as hosts for the three HVCV VLPs and for PBCV-1. Two of the zoochlorella isolates (NC64A and ATCC 30562) and, more recently, Nla, originally from United States isolates of P. bursaria, supported virus growth (181). The progeny particles were identical to PBCV-1, and they were infectious. This fulfilled Koch's postulates, and PBCV-1 is an authentic virus. More importantly, PBCV-1 formed plaques on lawns of Chlorella strain NC64A (Fig. 2), providing a sensitive bioassay for PBCV-1. None of the HVCV viruses replicated in the zoochlorellae from paramecia, and neither PBCV-1 nor HVCV replicated in any of ca. 50 additional free-living Chlorella or Chlamydomonas strains tested to date. The plaque assay for PBCV-1, together with the ability to synchronously infect the alga, allows one to study PBCV-1 by using bacteriophage technology. For example, one-step growth studies, life cycle studies, and genetic studies are feasible. The lack of a sexual stage in the host Chlorella organism and the current absence of an effective transformation system for the alga are disadvantages of the system. After developing the PBCV-1 plaque assay, we assayed freshwater samples collected throughout the United States and other parts of the world for plaque-forming viruses. As will be elaborated in the sections on properties of Chlorella strain NC64A viruses in addition to PBCV-1 and on the natural history of the Chlorella viruses, plaque-forming viruses on Chlorella strain NC64A are common in continental American, Chinese, and Japanese, but not European, fresh water. The viruses infecting Chlorella strain NC64A are referred to as NC64A viruses. Once we realized that NC64A viruses were common, we examined our original culture of paramecia for virus by the plaque assay. This culture, which had been stored at room temperature for about 18 months, still had viable green paramecia. However, no plaque-forming viruses were detected in the culture, nor could they be detected in zoochlorellae following isolation of the zoochlorellae from these paramecia (179). Consequently, the appearance of VLPs in the original, isolated paramecium zoochlorellae probably resulted from an external infection. Plaque-forming viruses which infect Chlorella strain SAG241-80 (69, 70) and Chlorella strain Pbi (referred to as Pbi viruses) (132, 134) have also been discovered in Soviet and European fresh water. Chlorella strains SAG-241-80 and Ebi are zoochlorellae originally cultured from European isolates of P. bursaria.
PROPERTIES OF CHLORELLA VIRUSES General Characteristics of PBCV-1 PBCV-1, the prototype NC64A virus, is easy to purify, and 80 to 105 A260 units (8 to 10 mg of PBCV-1) and 1.4 x 1012 to 2 x 1012 PFU of purified PBCV-1 are obtained per liter of lysate. Purified virus can be stored at 4°C for at least 1 year without detectable loss of infectivity. However, freezing inactivates PBCV-1.
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
593
B p
11
FIG. 1. Electron micrographs of the zoochlorellae present in H. viridis (A), immediately after isolation from the hydra (B), and after incubation of the isolated algae at 20°C for 6 h (C) and 20 h (D). Note the appearance of VLPs (arrow in panel C) in the algae at 6 and 20 h after isolation. Abbreviations: g, Golgi; n, nucleus; ch, chloroplast. The bar in C represents 1 ,um. Reproduced from reference 96 with permission from Academic Press, Inc.
594
MICROBIOL. REV.
VAN ETTEN ET AL.
1 week at 22°C in 5 mM EDTA or ethylene glycol-bis
lz--""ei'll-l.N I ...
il.
::::::.:---: :f:::::":"
...
f-
..
FIG. 2. Plaque assay of PBCV-1 on a lawn of Chlorella strain NC64A. The photograph was taken 4 days after the plate was prepared. However, plaques can be observed as early as 24 h after plating.
Electron micrographs indicate that PBCV-1 is a polyhedron with a multilaminate shell surrounding an electrondense core (Fig. 3). Frozen hydrated particles measure about 175 nm in diameter (Fig. 3A) (5). More recent electron micrographs indicate that PBCV-1 may have more substructure than was originally thought. For example, particles subjected to the quick-freeze, deep-etch procedure (53) contain flexible hairlike appendages (fibers) with swollen structures at the end; these appendages extend from at least some of the virus vertices (Fig. 3B). Micrographs of negatively stained particles also suggest that long fibers are attached to the outside of the capsid (small arrow in Fig. 3C). In addition, these micrographs suggest that some particles contain a distinctive 20- to 25-nm spike extending from one vertex (large arrow in Fig. 3C). The proportion of such particles suggests a unique vertex (50). DNA within the particle seems to be retracted from this vertex. The diameter of PBCV-1 is about 160 nm by comparison with lambda phage on the same grid (50). Speculation on the possible function of both the hairlike appendages and the spike structure will be mentioned in the section on virus attachment. It is obvious that additional research on the ultrastructure of PBCV-1 is required and that the particle may be more complex than a simple polyhedron. PBCV-1 sediments in sucrose density gradients at about 2,300S. The virus is disrupted on CsCl and Cs2SO4 density equilibrium gradients (187). The molecular weight of PBCV-1 has been estimated by two methods. From the DNA content (21 to 25%) and size (2.1 x 108 Da), the virion has a weight of 0.84 x 109 to 1 x 109 (184). Field flow fractionation gives a molecular weight of 1 x 109 (222). If one assumes a viral molecular weight of 1 x 109 and an extinction coefficient A 260 of 10.7 (uncorrected for light scattering), one A260 unit of PBCV-1 contains about 5.6 x 1010 virus particles (184). Experimentally we obtain 1.5 x 1010 to 3 x 1010 PFU per A260 unit of PBCV-1. Thus, 25 to 50% of the viral particles form plaques. PBCV-1 was treated with several reagents under a variety of conditions in an attempt to produce empty capsids and/or stable subviral core particles (156, 188). PBCV-1 is stable at pH 4 to 11. Virus infectivity is rapidly lost if PBCV-1 is stored above 45°C. PBCV-1 infectivity was stable for at least
(P-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA);
however, infectivity decreased slowly in 50 mM EDTA or EGTA. Virus treated with either 5 mM dithiothreitol or 5 mM dithioerythritol, but not 5 mM mercaptoethanol, rapidly lost infectivity. The dithiothreitol- or dithioerythritol-treated virus sedimented at the normal rate in sucrose gradients; however, electron micrographs of uranyl acetate-stained preparations revealed disrupted virus. To date, none of the treatments which decreased PBCV-1 infectivity have yielded either empty capsids or stable subviral core particles. Composition of PBCV-1 PBCV-1 contains about 64% protein, 21 to 25% dsDNA, and 5 to 10% lipid (156, 184). Analysis of PBCV-1 proteins by one-dimensional sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Fig. 4A) indicates more than 50 structural proteins ranging in apparent size from 10 to more than 200 kDa. Two-dimensional PAGE revealed additional proteins. However, some of the spots (especially the major capsid protein) smear on isofocusing gels, and repeated attempts to obtain "clean" two-dimensional patterns have been unsuccessful (157). One 54 kDa protein (Vp54) makes up about 40% of the total viral protein. If PBCV-1 is heated in denaturing buffer at 60°C, instead of 100°C, Vp54 migrates as a dimer with an apparent molecular weight of 104,000 (Fig. 4A, compare lanes 1 and 2). Vp54 is a glycoprotein. After deglycosylation of the protein with trifluoromethanesulfonic acid or hydrogen fluoride-pyridine, the protein migrates with an apparent size of about 49 kDa (196). The nature of the glycan linkage is unknown, although two observations suggest that it is 0 linked. Monensin, but not tunicamycin, inhibits both PBCV-1 synthesis and Vp54 synthesis (198). Incubation of Vp54 with the enzyme endo-,-N-acetylglucosaminidase, which cleaves N-linked sugars, does not affect its migration on SDS-gels. A second protein, Vp135, also stains with Schiff's reagent and is probably a glycoprotein. Vp135, Vp7l, Vp54, and Vpl4.5 are labeled by treating intact virus with 125I and are believed to be surface proteins (156). Serological experiments, described below, indicate that the glycan is on the virus surface. The resistance of PBCV-1 to several proteases is consistent with this interpretation. PBCV-1 proteins were also analyzed for myristylation and phosphorylation by infecting cells in the presence of [3H]myristic acid or 32P043- (128). Four proteins (Vp135, Vp55, Vp54, and Vp27.5) were specifically labeled by myristic acid. The myristic acid label was resistant to hydroxylamine (pH 8.5) and alkaline methanolysis, suggesting an amide linkage. Six PBCV-1 proteins (Vp6O, Vp45, Vp36, Vp29, Vp2O, and Vpl4) were labeled with 32P043- suggesting that they are phosphoproteins. The major lipids in PBCV-1 are phosphatidylcholine, phosphatidylethanolamine, and an unidentified component (156). PBCV-1 loses infectivity rapidly in chloroform and more slowly in ethyl ether or toluene; thus the lipid is required for infectivity. Sedimentation of organic solventtreated particles on sucrose gradients revealed that chloroform disrupted the virus. Ethyl ether treatment for 1 week did not alter the sedimentation properties of the virus. However, the ether-treated virus was swollen and partially disrupted after staining with uranyl acetate. PBCV-1 infectivity is unaffected by incubation with several detergents. For example, 2% Triton X-100, 2% Nonidet P-40, and 2% sodium deoxycholate have no effect on viral
* . e : . : *: : .Fse .s:i
*: A ..:. h. :.:S :.: SS: .::ee: {
i
.eSZ..Z.
*+S W: ;A: *\eS>{....;
':
:e
.: .:.
:e
..
..
..
.:
.:
.. ..
.::
.:.
..
.:
::
.: e.,
t-'.st>
.. *:
.;.
..
.:
_
..
.:
FIG. 3. Morphology of PBCV-1. (A) Frozen-hydrated particles (photograph kindly provided by Tim Baker). Note that the capsid appears as a double-layered structure in some of the particles. Bar, 200 nm. (B) PBCV-1 subjected to the quick-freeze, deep-etch procedure (53) (photograph kindly provided by John Heuser). Note the presence of flexible hairlike fibers, which appear swollen at their ends, on the surface of the particles. (C) PBCV-1 negatively stained with uranyl acetate (photograph kindly provided by Roger Hendrix). The smaller particles are lambda phage. Note that (i) long fibers are associated with the particles (small arrow), (ii) a distinctive 20- to 25-nm spike structure (large arrow) extends from one vertex of the particle, (iii) the particle appears to be packed asymmetrically, and (iv) an interior membrane appears to be located inside the capsid which extends to the unique vertex. 595
596
MICROBIOL. REV.
VAN ETTEN ET AL.
A
B 2
1 -w
BA
PST II
_MHI ID I H.
_l :.4
8268-
OR
0.
29-
V ,
_
4
.
dam "-a
a0wu
_e
17.2-
*FIG. 4. SDS-polyacrylamide gel of PBCV-1 proteins and PBCV-1 DNA restriction fragments. (A) Fluorogram of "4C-labeled PBCV-1 structural proteins following incubation at 60°C for 1 h (lane 1) or at 60°C for 55 min followed by 100°C for 5 min (lane 2). The material marked with an asterisk is presumed to be lipid. Note that the major viral protein, Vp54 (0), exists as a dimer in lane 1 and as a monomer in lane 2. Reproduced from reference 156 with permission from Academic Press, Inc. (B) PBCV-1 DNA restriction fragments: BamHIII (lane 1), HindIll (lane 2), and PstI (lane 3). Reproduced from reference 39 with permission from Martinus Nijhoff Publishers.
infectivity. However, PBCV-1 loses infectivity slowly in 2% sodium N-lauroyl sarcosine and rapidly in 2% SDS. These data, together with the ultrastructural data (i.e., an internal lipid layer is apparent in Fig. 3C), suggest that the lipid component is located inside the outer glycoprotein capsid. PBCV-1 Genome PBCV-1 contains a large dsDNA genome with a 40% G+C content (189). PBCV-1 DNA contains methylated bases. 5mC makes up 1.9% of the total cytosines, and 6mA makes up 1.5% of the total adenines. The size of the PBCV-1 genome, 333 kbp, was derived initially from summing restriction fragments. PBCV-1 DNA contains at least 41 BamHI fragments, 44 HindIIl fragments, and 50 PstI fragments (Fig. 4B) (39). Analysis of intact DNA by pulsed-field electrophoresis verified the size of PBCV-1 DNA at about 333 kbp (138). We initially reported that the PBCV-1 DNA restriction map was circular (39). This map was based on analyses of both cloned DNA and Southern transfers probed in some cases with gel-purified fragments that could not be cloned. From these experiments we concluded that PBCV-1 DNA was either a covalently closed circle, circularly permuted, or linear with terminal repetition. Subsequent experiments have established that the genome is a linear nonpermuted molecule and that the termini contain identical inverted
repeats. Furthermore, the termini have covalently closed hairpin ends (138, 167). PBCV-1 DNA terminal fragments were identified by incubating intact DNA with Bal31 exonuclease, which can initiate digestion in the single-stranded region of hairpin ends (see, e.g., references 6, 9, and 221), for various times before digestion with either HindIII, BamHI, or PstI restriction endonucleases (138). Two HindIII fragments (H4 and H17), two BamHI fragments (BlO and B15a), and two PstI fragments (P17 and P31) disappeared with increased Bal 31 exonuclease treatment, suggesting that these fragments were termini. Hybridization studies established that restriction fragments BlSa, H4, and P31 (arbitrarily called the left end) were at one terminus and that B10, H17, and P17 were at the other terminus. A revised restriction map beginning with Bl5a at zero and ending with B10 at 333 kbp is shown in Fig.
5. The terminal fragments, e.g., P31 and P17, hybridize with another, indicating terminal repetition. The termini (minus the hairpin) have recently been cloned and sequenced to within about 40 to 60 bases of the end (167). The termini contain identical inverted repeats of at least 2,185 bases, after which the sequence diverges. The inverted repeats contain two small open reading frames (ORFs) and several direct repeats. However, neither of the ORFs nor the remainder of the inverted repeats appear to be transcribed one
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
597
B31
B1 5a H4 P31
B2
I
I
I
P24
| |P14 P36a
0
|
H12a
H27b
|PS 10
B141 616 B8a HI8aI H24bI H23b I HIa P1O
20
IB2311
B12b
u
P13
B7b
I
I P9a 7080
P34b
Bib
I
H14
P4
|
110
H19 P15 I
50
Blb I P22
H3
Pi8c
I
100
Bla HI 8b
Hib
|
H29 130
P3b
P28b
B17 H13 90
B7c
|
I
H21 |
40
I H22bI
P26a 1,1 I P35 0 P38 P39
|HI8a |P25b
B32
II I
B14
H8b
P30 |P3a 30
P37
50
1 100
I
| H8c
B28
H3 P21
l'IB241
B8b
120
P32
I
P2c
140
150
B30
B1a H1b P2c
|
I
B12a
PI8b
i
150
619 |
|
|I H28
16
H9
P2a
170
160
B11
| B21|
H7
B4
H16
I
P28a
I
HUa
P33
IP34a4
P11
1;O
180
200 B25
84 HUa
|618b
-
P 1
|, I H24a| H27a
P8
200
1B26
63
| 620b
IH25 H23c
H20b
H15
PS
210
7a
||
H2 P27
220
P9b
|
P23
230
P25a
|
240
250
629 B7a H5c J P18a
B9 |
B5 Hil H
,H26| P29 PlBa
P16b
B6
|H20a
6B15b
300
320
B22 H22a
I
H5b
P16c
H17 P20
310
P7
B13
610
|H8a
P2b
H1O
H12b
P26b 280 P36b
270 | B20a
H5b
B18a H23a
P12
260
250
8B27
P17
330
333
FIG. 5. Revised PBCV-1 restriction map. The restriction sites for BamHI, HindlIl, and PstI are shown. Fragments Bl5a, H4, and P31 were placed at the left terminus, coordinate zero, and fragments B10, H17, and P17 were placed at the right terminus.
during PBCV-1 replication. The remainder of the PBCV-1 genome contains primarily unique DNA and has the potential to code for several hundred proteins (39). Serology of PBCV-1 Polyclonal antiserum prepared against intact PBCV-1 completely inhibits PBCV-1 infection by agglutinating the particles even at a dilution of 1:500. Western immunoblot analysis indicates that although the antiserum reacts with many of the virus proteins, it reacts most strongly with the major capsid protein Vp54 (197). If Vp54 is deglycosylated with trifluoromethanesulfonic acid or treated with periodic acid after transfer to nitrocellulose, the protein no longer reacts with PBCV-1 antiserum. Spontaneously derived antiserum-resistant mutants of PBCV-1 can be isolated at a frequency of about 10-6. Polyclonal antiserum was initially prepared against one of these mutants, named EPA-1. EPA-1 antiserum completely inhibits EPA-1 infection (dilution of 1:500), but has no effect (dilution of 1:10) on PBCV-1 infection (19). EPA-1 major capsid protein migrates slightly faster, with an apparent molecular mass of 53 kDa, than PBCV-1 Vp54. Like antiserum to PBCV-1, antiserum to EPA-1 reacts with the glycosylated but not the deglycosylated form of the major capsid protein on Western blot analysis. Subsequently, we isolated mutants resistant to both PBCV-1 and EPA-1 antiserum and prepared polyclonal
antiserum against some of these mutants (197). The antiserum-resistant mutants have several interesting properties. (i) Four PBCV-1 serotypes have been identified to date. All antisera reacted only with the virus which induced the antiserum. (ii) The major capsid proteins from all the mutants migrated slightly faster on acrylamide gels than did those of the strains from which they were derived. All major capsid proteins comigrated on SDS-gels after deglycosylation. (iii) Western blot analysis of the major capsid protein from the mutants, before and after deglycosylation, indicates that the serological differences among the viruses is due to a change in the carbohydrate moiety(ies) of the major capsid protein. (iv) The ratio of seven sugars (fucose, galactose, glucose, xylose, mannose, arabinose, and rhamnose) associated with the major capsid protein of PBCV-1 and the mutants varied in a predictable manner which was related to their serotype and the migration of the major capsid protein. (v) Each of the four serotypes arises from apparent single mutations in PBCV-1. This suggests that at least four virus gene products influence the PBCV-1 serotypes and that these genes are in a common pathway involved in glycosylation. Typically, carbohydrate moieties of virus glycoproteins are added in the rough endoplasmic reticulum and Golgi apparatus by host glycosyltransferases (142). Growing a virus in a different host (e.g., herpesvirus [77, 78], and influenza virus [26, 107]) can change the nature of the glycan.
598
VAN ETTEN ET AL.
Mutations that alter the polypeptide directly, e.g., changing a serine or asparagine to another amino acid (e.g., vesicular stomatitis virus [79]), can eliminate the glycan. We are presently testing the mutation possibility by cloning and sequencing the major capsid protein from PBCV-1 and some of the serological mutants. If the major capsid protein in PBCV-1 and the serotypes have identical amino acid sequences, this will confirm direct participation of viral enzymes in glycosylation.
Additional Chlorella Strain NC64A Viruses Hundreds of plaque-forming viruses, all of which infect Chlorella strain NC64A, have been isolated from fresh water collected throughout the continental United States and from China. Thirty-six of these viruses have been partially characterized and can be grouped into 16 classes (Table 2). Like PBCV-1, each of these viruses contains many structural proteins and a large dsDNA genome (at least 300 kbp) with 40% G+C, and all are chloroform sensitive. The viruses are polyhedra, with diameters of 150 to 190 nm. The DNAs of some hybridize extensively with PBCV-1 DNA, whereas others hybridize poorly (144, 227). The viruses can be distinguished from PBCV-1 and from each other by plaque size, antiserum sensitivity, DNA restriction patterns, sensitivity of the DNAs to restriction endonucleases, and the nature and abundance of methylated bases. DNA from each of the viruses contains 5mC in amounts varying from 0.12 to 47.5% of the total cytosine (Table 2). In addition, 24 of the 36 viral DNAs contain 6mA in amounts varying from 1.45 to 37% of the total adenine. The virus DNAs do not contain 4mC. Thus these viruses are a rich source of methylated DNAs. Such DNAs may be useful for studying the biological and physical properties of methylated DNAs. Growth curves revealed that viruses that form plaques similar in size to PBCV-1 (e.g., NC-1A and IL-3A) replicate at about the same rate as PBCV-1, whereas viruses that form small plaques (e.g., SC-1A, CA-4A, and NY-2A) take about two to three times longer to replicate (190). The burst size for each of the viruses is typically 200 to 350 PFU.
Viruses Infecting Chlorella Strain Pbi Plaque-forming viruses that infect an exsymbiotic Chlorella strain (Chlorella strain Pbi) of a European isolate of P. bursaria have also been isolated from European fresh water (132, 134). The Pbi viruses, like the NC64A viruses, are large polyhedra with diameters of 140 to 150 nm, are chloroform sensitive, have many structural proteins, and have large dsDNA genomes of at least 300 kbp that contain methylated bases (Table 3). However, the Pbi viruses are serologically distinct from the NC64A viruses, and their DNAs hybridize poorly with the NC64A virus DNAs. The Pbi virus DNAs have a higher G +C content (46%) than do the NC64A virus DNAs (40% G+C). Pbi viruses neither infect nor attach to Chlorella strain NC64A and the NC64A viruses neither infect nor attach to Chlorella strain Pbi (133). Additional viruses infecting Chlorella strain SAG-241-80 have been isolated and characterized from Soviet fresh water (69, 70). These viruses also infect Chlorella strain Pbi. Thus Chlorella strains Pbi and SAG-241-80 may be closely related or identicat.
MICROBIOL. REV.
REPLICATION OF PBCV-1
One-step growth experiments revealed that progeny PBCV-1 is first released about 3 to 4 h after infection and that virus release is complete by 8 to 10 h (Fig. 6A) (184). PBCV-1 release involves localized lysis of the cell wall. Thus one or more late virus gene products presumably degrade cell walls. Mechanical disruption of the cells releases infectious virus 30 to 50 min prior to spontaneous lysis. Consequently, infectious PBCV-1 is completely assembled inside the host and does not acquire lipids by budding through the host membrane. PBCV-1 also replicates in dark-grown Chlorella cells (Fig. 6B) or in light-grown cells treated with the photosynthetic inhibitor DCMU prior to virus infection. In both cases the time course of virus release is similar to that of untreated light-grown cells, and so PBCV-1 replication does not require host photosynthesis. However, the burst size, which is 200 to 350 PFU per cell in light-grown cells, is reduced about 50% in dark-grown cells. PBCV-1 infection rapidly inhibits host CO2 fixation. PBCV-1 replicates most efficiently in actively growing host cells and poorly in stationary-phase cells. Attachment of PBCV-l to Chlorella Strain NC64A PBCV-1 attaches rapidly and irreversibly to the external surface of cell walls, but not to protoplasts, of Chlorella strain NC64A with an adsorption rate of 5 x 10-9/ml/min. PBCV-1 attachment is specific since the virus attaches only to its host (93). Attachment occurs between pH 3.5 and 9 and between 5 and 25 mM MgCl2 or 5 and 100 mM NaCl (92). Each cell contains at least 5 x 104 binding sites, and Scatchard analysis indicates that 5,000 PBCV-1 particles can adsorb to each cell. Assuming that the alga is a sphere with a diameter of 5 ,Lm (94), approximately 2,500 virus particles of 175 nm would cover the surface of a single cell. Other estimates of the size of PBCV-1 have been smaller, 160 nm (50) or 139 nm (222), which means that more virus particles could fit on the surface of a single cell. From these data we concluded that (i) the entire surface of the Chlorella cell wall contains PBCV-1 receptors, (ii) individual receptor sites have similar affinities, and (iii) at most only a small percentage of the binding sites are used at any one time. The virus receptor is stable to heat and organic solvents since PBCV-1 attaches equally well to isolated walls of Chlorella strain NC64A even if the walls are first heated to 100°C or extracted with organic solvents, detergents, or high-salt solutions. The receptor is also resistant to many proteases, cellulase, and pectinase. However, exposure of the walls to HCl, pH 1, 0.5 M H2SO4, or 0.2 M NaOH for 1 h at 100°C inactivates the receptor (92). Treating walls with
the lectins wheat germ agglutinin, ricin, Lens culinaris lectin, or concanavalin A in the presence or absence of their specific hapten inhibitors also prevents virus attachment (91). This suggests that the lectins bind nonspecifically to the algal walls and that the lectin binding merely occludes viral attachment. With the exception of high concentrations (400 mM) of glucosamine and mannosamine, simple sugars such as glucose, N-acetylglucosamine, galactose, maltose, sucrose, fucose, rhamnose, arabinose, xylose, and N-acetylgalactosamine had no effect on PBCV-1 attachment (92). Attachment and infection of Chlorella strain NC64A by PBCV-1 is shown in Fig. 7A to E and in stereo views in Fig. 8. Attachment always occurs at a virus vertex. It is not known whether all virus vertices are identical and can serve as points of attachment. As mentioned above in the section
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
599
TABLE 2. Separation of Chlorella strain NC64A viruses into 16 classes' Class
1
Reaction with antibody tob: Restriction Plaque Level of methylation (mm) NY-2C Virus(m)gopMC group NY-2A 5mCd 6mAe PBCV-1 NYs-1 size
NE-8D NYb-1 CA-4B
3 3 3
Yes Yes Yes
No No No
No No No
2
AL-lA NY-2C NC-lD
3 3 3
No No No
Yes Yes Yes
3
PBCV-1 NC-lC
3 3
Yes Yes
4
CA-lA CA-2A IL-2A IL-2B IL-3A IL-3D
3 3 3 3 3 3
SC-lA SC-lB
6
7
NWf
ISV No
No
A A\
No No
A A
0.12
ND ND
Partial Partial Partial
Partial Partial Partial
A A A
0.45 0.39 0.33
ND ND ND
No No
No No
No No
C C
1.86 1.72
1.5 1.6
Yes Yes Yes Yes Yes Yes
No No No No No No
No No No No No No
No No No No No No
B B B B B B
1 1
Yes Yes
No No
No No
No No
D D
1.94 2.04
7.3 7.5
NC-lA
3
Yes
No
No
No
D
7.1
7.3
NE-8A AL-2C MA-lE NY-2F CA-iD NC-lB
3 3 3 3 3 3
Yes Yes Yes Yes Yes Yes
No No No No No No
No No No No No No
No No No No No No
E E E E E E
14.3
8.1
14.9 14.6
8.1 8.1
13.4
8.2
NYs-1
1
No
Partial
Yes
Yes
F
47.5
11.3
IL-5-2s1 AL-2A MA-iD NY-2B
1 1 1 1
No No No No
Partial Partial Partial Partial
Yes Yes Yes Yes
Yes Yes Yes Yes
G G G G
45.0 35.8 47.0 36.5
16.1 14.6 16.7 16.2
10
CA-4A
1
No
Partial
Yes
Yes
H
39.8
19.6
11
NY-2A
0.5
No
Partial
Yes
Yes
I
44.9
37.0
12
XZ-3A
1
Yes
No
No
No
J
12.8
2.2
13
SH-6A BJ-2C
1 1
Yes Yes
No No
No No
No No
K K
12.6
12.8
10.3 11.5
14
XZ-6E
1
Yes
No
No
No
E
21.2
15.2
15
XZ-4C
1
No
No
Yes
Yes
G
46.7
20.8
16
XZ-5C XZ-4A
1 1
No No
No No
Yes Yes
Yes Yes
H H
42.7 44.1
27.9 28.3
5
8 9
No
No
0-44 v rl 1.60 .
ND
10.0
ND
9.4 10.9 9.7 12.6
ND ND ND ND
a The separation is based on at least one of the following criteria: plaque size, reaction with four viral antisera, sensitivity of DNA to 13 restriction endonucleases, on percentage of 5mC and 6mA in genomic DNA. b A 5-,ug sample of purified virus was mixed with twofold serial dilutions of antiserum, and the precipitate was monitored 2 h later. Yes means that a reaction
was obtained at an antibody dilution of 1:64 or greater; Partial means a dilution of 1:8 to 1:32; and No means a dilution of 0 to 1:4. ' Viral DNAs were tested for susceptibility to the 13 restriction endonucleases BglII, EcoRI, BamHI, Sall, XhoI, Bcll, Pstl, HindIII, SstI, MboI, DpnI, MspI, and HapIl. The DNAs were grouped according to the resistance of the DNAs to these enzymes: group A, DpnI; group B, Hindlll, SstI, and Dpnl; group C, Bcll and MboI; group D, Sall, XhoI, BcIl, Pstl, and Mbol; group E, Sall, XhoI, BclI, Pstl, HindlIl, SstI, and MboI; group F, BamHI, BclI, PstI, HindIII, SstI, MboI, MspI and HpaII; group G, BamHI, Sall, XhoI, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII; group H, EcoRI, BamHI, Sail, XhoI, Bcll, PstI, HindlIl, SstI, MboI, MspI, and HpaII; group I, BglII, EcoRI, BamHI, Sall, Xhol, Bcll, PstI, HindIII, SstI, MboI, MpsI, and HpaII; group J, Bcll, HindIII, SstI, and MboI; group K, Bcll, PstI, HindIII, SstI, and MboI. d Percentage of 5mC per C plus 5mC plus deoxyuridine. e Percentage of 6mA per A plus 6mA plus deoxyinosine. f ND, not detected.
MICROBIOL. REV.
VAN ETTEN ET AL.
600
TABLE 3. Properties of five Chlorella strain Pbi viruseSa Plaque
Restriction
1 1 1 1 1
M
Virus
groupb
size (mm)
CVA-1 CVB-1 CVG-1 CVM-1
CVR-1
Level of methylation 5mCC 6mAd
NDe 17.7 ND 10.1 ND
43.1 42.7 19.2 41.9 14.2
N L F L
a None of these viruses reacted with polyclonal antisera to the Chlorella strain NC64A viruses. b Viral DNAs were tested for susceptibility to the 13 restriction endonucleases BglII, EcoRI, BamHI, Sall, XhoI, BMII, PstI, HindIII, SstI, MboI, DpnI, MspI, and HpaII. The DNAs were grouped according to the resistance of the DNAs to these enzymes: group F, BamHI, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII; group L, SalI and DpnI; group M, BamHI, Sall, Hindlll, SstI, DpnI, MspI, and HpaII; group N, EcoRI, BamHI, Sall, BclI, PstI, HindIII, SstI, MboI, MspI, and HpaII. c Percentage of 5mC per C plus 5mC plus deoxyuridine. d Percentage of 6mA per A plus 6mA plus deoxyinosine. e ND, none detected.
on general characteristics of PBCV-1, certain negatively stained preparations of PBCV-1 suggest that one vertex contains a spike structure which is consistent with a unique vertex. The stereo views (Fig. 8) indicate that the virus is attached to the wall by hairlike fibers which originate from at least some virus vertices. These interesting photographs suggest that the tips of the hairlike fibers are responsible for the initial recognition and attachment of the virus to the host. They could also be involved in orienting 4 unique virus vertex to the host surface. A scanning electron micrograph of numerous PBCV-1 particles attached to the host is shown in Fig. 9.
PBCV-1 attachment is followed by dissolution of the host wall at the point of attachment (Fig. 7B to D) and entry of the viral DNA and associated proteins into the cell, leaving an empty capsid on the host surface (Fig. 7E) (93). Since the virus also attaches to and digests Chlorella strain NC64A wall fragments which have previously been heated to 100°C, the virus carries the hydrolytic enzyme(s) (Fig. 7F). The cell-wall-degrading activity is located inside the virus particle (possibly in the spike structure), because (i) intact, protease-treated PBCV-1 particles still attach and digest a hole in the host wall and (ii) cell-wall-digesting activity can be isolated from purified, osmotically disrupted PBCV-1 particles (20). This suggests that attachment might trigger a change in the virus structure. However, attachment to and digestion of wall fragments does not lead to empty PBCV-1 capsids; thus DNA release appears to require a host function(s) (e.g., perhaps the virus internal membrane fuses with the host membrane). Attempts to release DNA from PBCV-1 attached to isolated walls by adding various agents and cell fractions have so far been unsuccessful (179).
Intracellular Site of PBCV-1 Replication The destination of the infecting DNA and the intracellular site of PBCV-1 transcription and DNA replication are unknown. PBCV-1 proteins are synthesized on cytoplasmic ribosomes and not organelle ribosomes, since cycloheximide, but not chloramphenicol, inhibits viral replication (156). PBCV-1 replication does not require host transcription, since PBCV-1 replicates in UV-irradiated cells (182). Such cells are unable to form colonies, and endogenous host RNA and DNA syntheses are reduced to background levels. However, PBCV-1 replication is slow (latent period of 16 to 20 h) and the burst size is small (2 to 10 PFU per cell) in 105 -B
A 0
, -
--
-----_
-0
/1 I
0
Ji
2
104
0
1
m I
LL I
0
-i
q
2
II
G&
IL
11
03
IIp ~0
1o2
02 100
200
300
400
TlblE, MIN.
500
600 101 0
100
200
300
400
500
600
TIME. MIN.
FIG. 6. One-step growth curves of PBCV-1 on Chlorella strain NC64A grown in the light (A) or in the dark (B). Symbols: lysis; 0, mechanically disrupted algae. Reproduced from reference 184 with permission from Academic Press, Inc.
0,
spontaneous
1~~~ ~~e>6 f
A7~~~~
A. I 1A
I..,
r
s.iF.
t... ,.,.
-jt-
0.
or
.
?b
..
Pt.i
. ;rg e
I
-.
fli , sf'.
.,
½~~~~~~~~~4-~
x
'.
-.
*~~~~~~~v I.,
D
C
,W: -s >f -I "m:o;f
1:
.L
wy
,.
:,
*< 9 'i ;_ q
rI
FIG. 7. Infection of Chiorella strain NC64A by PBCV-i. (A) Viral particle in close proximity to the alga. (B and C) Attachment of PBC Vto the algal wall and digestion of the wall at the point of attachment. (D) Viral DNA beginning to enter the host. (E) An empty viral capsid remaining on the surface of the host. (F) PBC V-i attachment and dissolution of a Chiorella cell wall fragment. Note that (i) viral attachment always occurs on the external side of the wall (i.e., the internal side of the wall curls inward) and (ii) DNA is not released from viral particles attached to wall fragments. The size markers in panels E and F represent 100 nm and 200 nm, respectively. Reproduced from reference 93 with permission from Academic Press, Inc.
601
602
VAN ETTEN ET AL.
MICROBIOL. REV.
FIG. 8. Stereo views of PBCV-i attached to Chlorella strain NC64A. The cells were quick-frozen and deep-etched. (A) Side view of the particles attached to the cell wall; (B) top view of the particles attached to the cell wall. Note that the particles are attached to the wall via hairlike fibers, which appear to originate from the vertices of the virus. Photographs kindly provided by John Heuser.
UV-treated cells. PBCV-1 replication also does not require labile host factors, since the virus replicated in UV-irradiated cells that had been incubated in the dark for up to 8 h before infection. The host cells change cytologically beginning about 1 to 2 h after infection (94). In the nucleus, the nucleolus disintegrates and chromatin heterochromicity increases. The nucleus, mitochondria, and Golgi apparatus become appressed to the chloroplast, leaving one or more finely granulated electron-translucent areas in the cytoplasm (labeled V in Fig. 1OA to D). Since PBCV-1 ultimately forms in these areas, we have designated these areas virus assembly centers. Early stages of PBCV-1 morphogenesis in the virus assem-
bly centers are seen in a few cells by 1 h postinfection (p.i.) and are common by 2 h p.i. (Fig. 10 A and B). Short membranous structures, which are vesicular or angular in shape (arrow in Fig. 10 B and C), first appear in the virus assembly centers. These membranes appear to function as templates for capsid assembly. More dense material (presumably protein) appears on the exterior surface of these membranes, giving rise to angular configurations (Fig. 1OA and B). These developing capsids are bounded by a distinct electron-dense layer; a thicker translucent region is visible directly underneath this dense outer layer (Fig. lOC and D). Capsids are usually formed at the periphery of the virus assembly centers, producing the rosette pattern seen in Fig.
VIRUSES OF EUKARYOTIC ALGAE
VOL. 55, 1991
603
FIG. 8-Continued.
lOB. Apparent complete capsids are assembled prior to DNA accumulation. A ring of virus particles surrounding a virus assembly center, which are in various stages of packaging DNA, are seen in Fig. lOD. These observations suggest that PBCV-1 probably packages DNA through an opening in the capsid. By 4 to 5 h p.i. the cells contain numerous filled virus particles, which are distributed throughout the cytoplasm. Sometimes a nuclear membrane is still visible at late stages of infection (Fig. 10E). Mitochondria and the chloroplast disintegrate in late stages of PBCV-1 replication. So far there is no evidence for nuclear or organellar roles in viral replication. If the nucleus is not involved, transcriptionassociated enzymes, such as RNA polymerase(s) and pro-
cessing enzymes, must be imported into the cell with the infecting DNA. PBCV-1 has not been effectively assayed for these enzymes. We have been unsuccessful in brief attempts to detect RNA polymerase activity either in intact PBCV-1 or in PBCV-1 that had been disrupted osmotically or by freezethawing (15). Although PBCV-1 may not contain a functional RNA polymerase, we suspect that our failure may reflect unsatisfactory disruption methods. We predict that PBCV-1 imports at least some of the enzymes involved in its replication. Since PBCV-1 is assembled in specific regions in the cytoplasm, the cytoskeleton might be expected to actively target virus proteins, including the glycoproteins, to the
604
VAN ETTEN ET AL.
MICROBIOL. REV.
Although recycling from host DNA may supply some of these intermediates, it can supply only 10 to 25% of the requirement. Consequently, virus-infected cells may be an excellent model system for studying nucleotide metabolism. One obvious test for recycling is to label host DNA prior to infection and determine whether the label accumulates in virus DNA. Unfortunately, Chlorella strain NC64A lacks thymidine kinase (180) and so host DNA cannot be labeled
FIG. 9. Scanning electron micrograph of PBCV-1 particles attached to Chlorella strain NC64A (photograph kindly provided by Kit Lee). Bar, 1 plm.
assembly center. However, several cytoskeleton-disrupting agents, which inhibit either tubulin or actin functions, had no effect on PBCV-1 replication even at concentrations much higher than those required to inhibit growth of the host Chlorella cell (112). Effect of PBCV-1 Infection on Host DNA, RNA, and Protein Syntheses
PBCV-1 infection dramatically changes host macromolecular syntheses. The following changes occur in DNA metabolism (180). (i) Virus infection almost immediately inhibits host DNA synthesis. (ii) PBCV-1 DNA synthesis begins 50 to 60 min after infection and requires de novo synthesis of a protein(s). This implies that PBCV-1 directs the synthesis of at least some of its own DNA synthetic machinery. New DNA polymerase activities appear in infected cells (139, 140). (iii) PBCV-1 infection leads to the degradation of host chloroplast and nuclear DNAs within 1 to 2 h p.i. (Fig. 11). Presumably a virus-encoded protein(s) mediates this response since cycloheximide added at the time of infection prevents host DNA degradation. Host nuclear and chloroplast DNA levels also decrease, albeit more slowly, if UV-inactivated algae are inoculated with PBCV-1 (209). (iv) The total DNA level in the host cell increases 3- to 10-fold by 5 h after PBCV-1 infection. Thus PBCV-1 replication requires a large increase in deoxynucleotide intermediates.
specifically. PBCV-1 infection also alters RNA metabolism (145). (i) Viral infection rapidly inhibits host RNA synthesis. (ii) Chloroplast rRNAs, but not cytoplasmic rRNAs, are degraded beginning about 30 min p.i. Presumably a virusencoded protein(s) contributes to this process, since cycloheximide added at the time of PBCV-1 infection prevents chloroplast rRNA degradation. (iii) PBCV-1 transcription is programmed, and early transcripts appear within 5 to 10 min after infection. (iv) Some, but not all, early viral transcripts are synthesized in the absence of de novo protein synthesis. The synthesis of later transcripts requires translation of an early gene product(s). (v) Late viral transcription begins about 1 h p.i., at the same time viral DNA synthesis begins (Fig. 12). (vi) A few early transcripts continue to be synthesized after virus DNA synthesis begins (45). Little is known about protein synthesis following virus infection. PBCV-1 infection rapidly inhibits host protein synthesis by at least 75%. We have briefly investigated virus protein synthesis in PBCV-1-infected cells by examining proteins on polyacrylamide gels (215). These experiments were less than satisfactory because (i) the synthesis of some host proteins continued (at least briefly) after infection and (ii) the specific labeling of viral proteins was low, even after long labeling periods (more than 1 h) with high-specificactivity amino acids. This latter finding suggests that PBCV-1 infection may expand amino acid pools. The increase could reflect recycling from degraded host protein or from increased amino acid synthesis. A complicating factor is that PBCV-1 has the potential to code for several hundred proteins, which would require two-dimensional gels for adequate resolution. TRANSCRIPTION OF PBCV-1 As mentioned in the preceding section, PBCV-1 transcription can be conveniently divided into early and late stages. The junction between these stages is about 1 h p.i., which coincides with the start of virus DNA synthesis. Interestingly, the incorporation of [3H]adenine into polyadenylatecontaining RNA also changes abruptly at about 1 h after PBCV-1 infection. Polyadenylate segments are virtually absent from late transcripts, whereas at least some early virus transcripts are probably polyadenylated (190). RNA isolated at various times after PBCV-1 infection was hybridized with each of four cloned fragments of PBCV-1 DNA (Fig. 12) (145). Both early and late genes are dispersed within the PBCV-1 genome, since each of the four cloned fragments hybridized to both early and late transcripts.
FIG. 10. Chlorella cells at 2 h (A to C), 5 h (D), and 6 h (E) after PBCV-1 infection. Parts of newly forming capsids (arrow in panel A) were located in defined areas in the cytoplasm (labeled v for virus assembly center in panels A to D) by 2 h p.i. Membranous materials which assume vesicular or angular shapes were usually seen in the virus assembly centers (arrow in panels B and C). The newly forming capsids can have a rosette appearance, which surrounds the virus assembly center (panel B). By 5 h p.i. (panel D) the cells contained a mixture of filled and partially filled capsids, empty capsids, and incomplete capsids. By 6 h p.i. (panel E), the cells contained many filled capsids which were distributed throughout the cytoplasm. Occasionally, cells contained aberrant virus capsids (F). Bars: 500 nm (panels A to D and F) and 1,000 nm (panel E). Reproduced from reference 94 with permission from Academic Press, Inc.
VOL. 55, 1991
~;Se$,g.A wsja35ve~ ~
VIRUSES OF EUKARYOTIC ALGAE
,,, ~ ~~~~~~~~~~~~~~~i.
i ~~~~~ N
;
5 a fi
X ;
WS2Atr 6
&
4~~~~~~~~~~~~~~~~~~~~~~1
I'
* r ve
.,... , ; ;g ,.,. ,kx !'
. , ,,:VA 3; $~I
..'r
, . . .4
AUZA
8K,K-
* A i ; ' . ?
i * '
TV;
ov
|:s;: ~~~'
eAW ^
..
;
* tj sw !Sg O
y S * * * ' + t s L " iryo4~~ ~ ~ ~ ~ ~ ~ N
i
4
w < ? >rRvntf > N.
f .Ssrv
1pm
b
V /
PIPI.e 1w,
.f
i
"Ok
C
-
0W-5).5um
_
0
d ~~~~~~~~~~~~~~~~~~~~~~~I _
_-
MICROBIOL. REV.
0. -- m
FIG. 15. Micrographs of the VLP from U. gigas. (a) Thin section of an infected cell containing hexagonal and pentagonal particles with two internal components in some particles. (b) Tailed and tailless VLPs negatively stained in 2%b phosphotungstic acid. (c) Tailed particle with a hexagonal capsid. Note the characteristic swelling in the middle of the tail. (d) Tailed particle with a swollen spherical capsid. Reproduced from reference 29 with permission from Academic Press, Inc.
VOL. 55, 1991
VIRUSES OF EUKARYOTIC ALGAE
615
nia VLPs. The Ectocarpus particles are released into the medium when host cells lyse, and they can infect healthy zoospores. The infected zoospores become paralyzed and settle to the bottom of the container, where they develop into plants that produce viruses in their sporangia. Expression of the Ectocarpus virus is temperature sensitive and is linked to differentiation of reproductive cells of the host. Notably, both research groups have preliminary evidence suggesting that the Feldmannia and Ectocarpus virus DNAs are integrated into their host DNAs (51, 104).
CONCLUDING COMMENTS
FIG. 16. VLPs near and attached to the surface of the marine nanoflagellate M. pusilla 15 min after inoculation. Unlike most eukaryotic algae, M. pusilla lacks a cell wall. Reproduced from reference 87 with permission from Macmillan Magazines Ltd.
sporangia (52), suggesting that VLP replication might be triggered by meiosis. Muller et al. (103) found viruses in Ectocarpus siliculosus, a related ectocarpoid species also isolated near the coast of New Zealand, which contained particles about 130 nm in diameter. These particles were located exclusively in the gametangia. The particles contain a 350-kbp linear dsDNA, which is about twice the size of the DNA from the Feldman-
Algal viruses, especially the Chlorella viruses, are interesting for several reasons and deserve investigation. The Chlorella viruses are examples of a new virus-host relationship, and their bacteriophagelike biology makes them excellent models for studying gene regulation and expression in photosynthetic eukaryotes. The inability to perform host genetics and to introduce foreign genes into the viruses remains a disadvantage. The genomes of these viruses are potential sources of control elements such as origin(s) of replication or promoters for plant genetic engineering. In fact, the upstream region of one of the Chlorella virus DNA methyltransferase genes (M.CviBIII) functions well as a promoter in both Agrobacterium tumefaciens and higher plants (100). Methylation levels of the Chlorella viral DNAs vary considerably; thus, some of these DNAs may be useful for molecular and biological studies on the effect of methylation on DNA structure and function. For example, it would be interesting to determine whether DNA-binding proteins
FIG. 17. Virus particles in Ectocarpus siliculosus (photograph kindly provided by Eric Henry). The particles are present only in the
sporangia.
616
VAN ETTEN ET AL.
of the viruses are adapted to levels of methylation. The Chlorella viruses contain several genes of commercial interest such as DNA methyltransferases and DNA site-specific endonucleases, some of which have unique specificities. In addition, the viruses contain host-wall-degrading enzymes which could potentially digest plant cell walls. Finally, preliminary results suggest that the viruses may contain glycosyltransferase genes. The common association of VLPs with zoochlorellae isolated from hydras and paramecia suggests that these particles may influence symbiosis. Viruses that infect Chlorella strain NC64A are common in fresh water; however, nothing is known about their potential ecological impact on the algal community. Although investigators are just beginning to look for viruses of other eukaryotic algae, it is already obvious that viruses are a major factor in marine algal ecology (153, 169). Besides limiting algal growth, the viruses could potentially mediate genetic exchange among native algae. Pearson and Norris (122) and Hoffman and Stanker (56) have also suggested that algae could serve as reservoirs or vectors for transmitting viruses to other forms of aquatic life, including animals. In this regard the Chlorella viruses have some similarities to important disease viruses such as the poxviruses and ASFV. Finally, at least some of these viruses, such as those infecting brown algae, appear to be expressed only in meiotic cells, suggesting highly specific coordination between the virus and the host life cycle. We encourage more investigators to work on these virus systems. ACKNOWLEDGMENTS We (J.V.E. and R.H.M.) thank the many technicians, undergraduate students, graduate students, postdoctoral associates, and visiting scientists who have worked in our laboratories over the past 10 years on the Chlorella viruses. In addition, many people at other institutions have graciously provided information and help on specific aspects of the viruses, as well as providing some of the photographs used in this paper. Special thanks go to Dwight Burbank, who has worked with the viruses since their discovery, and Kit Lee, who has taken many of the electron micrographs of the Chlorella viruses. We also acknowledge the many stimulating discussions with Myron Brakke over the years. A special acknowledgment goes to Malcolm Brown, who first pointed out the virus particles in one of our (R.H.M.) original electron micrographs. Finally, we thank Myron Brakke, Mike Nelson, Roy French, Reingard Grabherr, Yanping Zhang, Mike Groves, Eric Henry, Kyle Hoogland, and Jim Rosowski for reading and making many helpful suggestions on various aspects of the manuscript. Research from our laboratories has been supported by grants from the Department of Energy (J.V.E.), the National Institutes of Health (J.V.E.), the National Science Foundation (R.H.M.), the U.S. Department of Agriculture (R.H.M.), and the Solar Energy Research Institute (R.H.M.).
REFERENCES 1. Aach, H. G., S. Bortsch, and V. Feyen. 1978. Studies on Chlorella protoplasts: Demonstration of the protoplastic nature and the regeneration of the cell wall. Planta 139:257-260. 2. Adams, M. H. 1959. Bacteriophages. Interscience Publications, Inc., New York. 3. Atkinson, A. W., B. E. S. Gunning, and P. C. L. John. 1972. Sporopollenin in the cell wall of Chlorella and other algae: ultrastructure, chemistry and incorporation of "4C-acetate, studied in synchronous cultures. Planta 107:1-32. 4. Baker, J. R. J., and L. V. Evans. 1973. The ship fouling alga Ectocarpus. I. Ultrastructure and cytochemistry of plurilocular reproductive stages. Protoplasma 77:1-13. 5. Baker, T. S. Personal communication. 6. Barbour, A. G., and C. F. Garon. 1987. Linear plasmids of the bacterium Borrelia burgdorferi have covalently closed
MICROBIOL. REV.
ends. Science 237:409-411. 7. Berliner, M. D. 1977. Protoplast induction in Chlorella vulgaris. Plant Sci. Lett. 9:201-204. 8. Bestor, T., A. Laudano, R. Mattaliano, and V. Ingram. 1988. Cloning and sequencing of a cDNA encoding DNA methyltransferase of mouse cells: the carboxylterminal domain of the mammalian enzymes is related to bacterial restriction methyltransferases. J. Mol. Biol. 203:971-983. 9. Blackburn, E. H., and P. B. Challoner. 1984. Identification of a telomeric DNA sequence in Trypanosoma brucei. Cell 36:447457. 10. Blasco, R., I. D. L. Vega, F. Almazan, M. Aguero, and E. Vinuela. 1989. Genetic variation of African swine fever virus: variable regions near the ends of the viral DNA. Virology 173:251-257. 11. Bonnefoy, A. M., X. Kolenkine, and C. Vago. 1972. Particules d'allure virale chez les Hydres. C.R. Acad. Sci. (Paris) 275: 2163-2165. 12. Borowitzka, M. A., and L. J. Borowitzka (ed.). 1988. Microalgal biotechnology. Cambridge University Press, Cambridge. 13. Brooks, J., and G. Shaw. 1978. Sporopollenin: a review of its chemistry, palaeochemistry, and geochemistry. Grana 17:9197. 14. Brunt, A. A., and K. E. Richards. 1989. Biology and molecular biology of the furoviruses. Adv. Virus Res. 36:1-32. 15. Burbank, D. E., R. Grabherr, and J. L. Van Etten. Unpublished results. 16. Burbank, D. E., S. L. Shields, A. M. Schuster, and J. L. Van Etten. 1990. 5-Azacytidine resistant mutants of Chlorella virus IL-3A. Virology 176:311-315. 17. Cernichiari, E., L. Muscatine, and D. C. Smith. 1969. Maltose excretion by the symbiotic algae of Hydra viridis. Proc. R. Soc. London Ser. B 173:557-576. 18. Chapman, R. L., and N. J. Lang. 1973. Virus-like particles and nuclear inclusions in the red alga Porphyridium purpureum (Bory) Drew et Ross. J. Phycol. 9:117-122. 19. Chase, T. E., J. A. Nelson, D. E. Burbank, and J. L. Van Etten. 1989. Mutual exclusion occurs in a Chlorella-like green alga inoculated with two viruses. J. Gen. Virol. 70:1829-1836. 20. Ciuffetti, L., Q. Que, J. L. Van Etten, and R. H. Meints. Unpublished results. 21. Clitheroe, S. B., and L. V. Evans. 1974. Viruslike particles in the brown alga Ectocarpus. J. Ultrastruct. Res. 49:211-217. 22. Cock, J. M., L. L. Roof, N. F. Bascomb, C. W. Gehrke, K. C. Kuo, and R. R. Schmidt. 1990. Restriction enzyme analysis and cloning of high molecular weight genomic DNA isolated from Chlorella sorokiniana (Chlorophyta). J. Phycol, 26:361-367. 23. Cole, A., J. A. Dodds, and R. I. Hamilton. 1980. Purification and some properties of a double-stranded DNA containing virus-like particle from Uronema gigas, a filamentous eucaryotic green alga. Virology 100:166-174. 24. Danko, S., and J. L. Van Etten. Unpublished results. 25. DaSilva, E. J., and H. G. Gyllenberg. 1972. A taxonomic treatment of the genus Chlorella by the technique of continuous classification. Arch. Mikrobiol. 87:99-117. 26. Deom, C. M., and I. T. Schulze. 1985. Oligosaccharide composition of an influenza virus hemagglutinin with host-determined binding properties. J. Biol. Chem. 260:14771-14774. 27. Dodds, J. A. 1979. Viruses of marine algae. Experientia 35: 440 442. 28. Dodds, J. A. 1983. New viruses of eukaryotic algae and protozoa, p. 177-188. In R. E. F. Matthews (ed.), A critical appraisal of viral taxonomy. CRC Press, Inc., Boca Raton, Fla. 29. Dodds, J. A., and A. Cole. 1980. Microscopy and biology of Uronema gigas, a filamentous eukaryotic green alga, and its associated tailed virus-like particle. Virology 100:156-165. 30. Doermann, A. H. 1983. Introduction to the early years of bacteriophage T4, p. 1-7. In C. K. Mathews, E. M. Kutter, G. Mosig, and P. B. Berget (ed.), Bacteriophage T4. American Society for Microbiology, Washington, D.C. 31. Douglas, A. E., and V. A. R. Huss. 1986. On the characteristics and taxonomic position of symbiotic Chlorella. Arch. Microbiol. 145:80-84.
VOL. 55, 1991
32. Dunn, K. 1987. Growth of endosymbiotic algae in the green hydra, Hydra viridissima. J. Cell Sci. 88:571-578. 33. Eisenstadt, E. 1971. Transfer of photosynthetic products from symbiotic algae to animal tissue in Chlorohydra viridissima, p. 202-208. In H. M. Lenhoff, L. Muscatine, and L. V. Davis (ed.), Experimental coelenterate biology. University of Hawaii Press, Honolulu. 34. Franca, S. 1976. On the presence of virus-like particles in the dinoflagellate Gyrodinium resplendens (Hulbert). Protistologica 12:425-430. 35. Garon, C. F., E. Barbosa, and B. Moss. 1978. Visualization of an inverted terminal repetition in vaccinia virus DNA. Proc. Natl. Acad. Sci. USA 75:4863-4867. 36. Geshelin, P., and K. I. Berns. 1974. Characterization and localization of the naturally occurring cross-links in vaccinia virus DNA. J. Mol. Biol. 88:785-796. 37. Gibbs, A. Personal communication. 38. Gibbs, A., A. H. Skotnicki, J. E. Gardiner, E. S. Walker, and M. Hollings. 1975. A tobamovirus of a green alga. Virology 64:571-574. 39. Girton, L. E., and J. L. Van Etten. 1987. Restriction site map of the Chlorella virus PBCV-1 genome. Plant Mol. Biol. 9:245-257. 40. Goff, L. J. (ed.). 1983. Algal symbiosis: a continuum of interaction strategies. Cambridge University Press, Cambridge. 41. Gonzalez, A., A. Talavera, J. M. Almendral, and E. Vinuela. 1986. Hairpin loop structure of African swine fever virus DNA. Nucleic Acids Res. 14:6835-6844. 42. Goohra, R., and A. Granoff. 1979. Icosahedral cytoplasmic deoxyriboviruses, p. 347-399. In H. Fraenkel-Conrat and R. R. Wagner (ed.), Comprehensive virology, vol. 14. Plenum Press, New York. 43. Goorha, R., and K. G. Murti. 1982. The genome of frog virus 3, an animal DNA virus, is circularly permuted and terminally redundant. Proc. Natl. Acad. Sci. USA 79:248-252. 44. Goorha, R., D. B. Willis, and A. Granoff. 1977. Macromolecular synthesis in cells infected by frog virus 3. VI. Frog virus 3 replication is dependent on the cell nucleus. J. Virol. 21:802805. 45. Graves, M., and R. H. Meints. Unpublished results. 46. Gromov, B. V., and K. A. Mamkaeva. 1979. Phage-form infectious virus of the green alga Chlorococcum. Bull. Acad. Sci. USSR Biol. N2:181-186. 47. Gromov, B. V., and K. A. Mamkaeva. 1981. A virus infection in the synchronized population of the Chlorococcum minutum zoospores. Arch. Hydrobiol. Suppl. 60:252-259. 48. Hayes, W. 1968. The genetics of bacteria and their viruses, 2nd ed. John Wiley and Sons, Inc., New York. 49. Hellmann, V., and E. Kessler. 1974. Physiologische und biochemische Beitrage zur Taxonomie der gattung Chlorella. VIII. Die Basenzusammensetzung der DNS. Arch. Microbiol. 95:311-318. 50. Hendrix, R. Personal communication. 51. Henry, E., and R. H. Meints. Unpublished results 52. Henry, E. C., S. Lanka, and R. H. Meints. 1989. A stable virus infection in a brown alga. Abstr. XIII Int. Seaweed Symp., p. A98. 53. Heuser, J. 1980. Three-dimensional visualization of coated
vesicle formation in fibroblasts. J. Cell Biol. 84:560-583. 54. Hoffman, L. Personal communication. 55. Hoffman, L. R. 1978. Virus-like particles in Hydrurus (Chrysophyceae). J. Phycol. 14:110-114. 56. Hoffman, L. R., and L. H. Stanker. 1976. Virus-like particles in the green alga Cylindrocapsa. Can. J. Bot. 54:2827-2841. 57. Houts, G. E., M. Gravell, and A. Granoff. 1974. Electron microscopic observations on early events of frog virus 3 replication. Virology 58:589-594. 58. Huss, V. A. R., G. Huss, and E. Kessler. 1989. Deoxyribonucleic acid reassociation and inter species relationships of the genus Chlorella (Chlorophyceae). Plant Syst. Evol. 168:71-82. 59. Jolley, E., and D. C. Smith. 1978. The green hydra symbiosis. I. Isolation, culture, and characteristics of the Chlorella sym-
VIRUSES OF EUKARYOTIC ALGAE
617
biont of European Hydra viridis. New Phytol. 81:637-645. 60. Jones, P. A. 1984. Gene activation by 5-azacytidine, p. 165-187. In A. Razin, H. Cedar, and A. D. Riggs (ed.), DNA methylation: biochemistry and biological significance. Springer-Verlag, KG, Berlin. 61. Karakashian, M. W. 1975. Symbiosis in Paramecium bursaria. Symp. Soc. Exp. Biol. 29:145-173. 62. Karakashian, S. J., and M. W. Karakashian. 1965. Evolution and symbiosis in the genus Chlorella and related algae. Evolution 19:368-377. 63. Kawakami, H., and N. Kawakami. 1978. Behavior of a virus in a symbiotic system, Paramecium bursaria-zoochlorella. J. Protozool. 25:217-225. 64. Kelly, D. C. 1975. Frog virus 3 replication: electron microscope observations on the sequence of infection in chick embryo fibroblasts. J. Gen. Virol. 26:71-86. 65. Kessler, E. 1982. Chemotaxonomy in the Chlorococcales. Prog. Phycol. Res. 1:111-135. 66. Kessler, E., V. A. R. Huss, and M. Rahat. 1988. Speciesspecific ability of Chlorella strains (Chlorophyceae) to form stable symbioses with Hydra viridis. Plant Syst. Evol. 160:241246. 67. Klimasauskas, S., A. Timinskas, S. Menkevicius, D. Butkiene, V. Butkus, and A. Janulaitis. 1989. Sequence motifs characteristic of DNA [cytosine-N4] methyltransferases: similarity to adenine and cytosine-C5 DNA-methylases. Nucleic Acids Res. 17:9823-9832. 68. Koltin, Y., and M. J. Leibowitz (ed.). 1988. Viruses of fungi and simple eukaryotes. Marcel Dekker, Inc., New York. 69. Kvitko, K., and B. V. Gromov. 1984. New findings of a titratable infectious zoochlorella virus. Dokl. Akad. Nauk. SSSR 279:998-999. 70. Kvitko, K. V., I. I. Skoblo, E. S. Kraeva, E. V. Skovorodina, and T. A. Bulkina. 1985. Specificity of host range of zoochlorella viruses from Paramecium bursaria. All Union Meet. Microbiol. Soc. USSR 5:17. 71. LaClaire, J. W., and J. A. West. 1977. Virus-like particles in the brown alga Streblonema. Protoplasma 93:127-130. 71a.Lauster, R. 1989. Evolution of type II DNA methyltransferases: a gene duplication model. J. Mol. Biol. 206:313321. 72. Lauster, R., T. A. Trautner, and M. Noyer-Weidner. 1989. Cytosine-specific type II DNA methyltransferases: a conserved enzyme core with variable target-recognizing domains. J. Mol. Biol. 206:305-312. 73. Lee, R. E. 1971. Systemic viral material in the cells of the freshwater red alga Sirodotia tenuissima (Holden) Skuja. J. Cell Sci. 8:623-631. 74. Lemke, P. A. 1976. Viruses of eucaryotic microorganisms. Annu. Rev. Microbiol. 30:105-145. 75. Loefer, J. B. 1936. Isolation and growth characteristics of the zoochlorella of Paramecium bursaria. Am. Nat. 70:184-188. 76. Loos, E., and D. Meindl. 1982. Composition of the cell wall of Chlorella fusca. Planta 156:270-273. 77. Lundstrom, M., S. Jeansson, and S. Olofsson. 1987. Host cell-induced differences in the 0-glycosylation of herpes simplex virus gC-1. II. Demonstration of cell-specific galactosyltransferase essential for formation of 0-linked oligosaccharides. Virology 161:395-402. 78. Lundstrom, M., S. Olofsson, S. Jeansson, E. Lycke, R. Datema, and J. E. Mansson. 1987. Host cell-induced differences in 0-glycosylation of herpes simplex virus gC-1. I. Structures of nonsialylated HPA- and PNA-binding carbohydrates. Virology 161:385-394. 79. Luo, L., Y. Li, R. M. Snyder, and R. R. Wagner. 1990. Spontaneous mutations leading to antigenic variations in the glycoproteins of vesicular stomatitis virus field isolates. Virology 174:70-78. 80. Luria, S. E., J. E. Darnell, D. Baltimore, and A. Campbell. 1978. General virology. John Wiley & Sons, Inc., New York 81. Mann, M. B., R. N. Rao, and H. 0. Smith. 1978. Cloning of restriction and modification genes in E. coli: the HhaII system from Haemophilus haemolyticus. Gene 3:97-112.
618
MICROBIOL. REV.
VAN ETTEN ET AL.
82. Manton, I., and B. S. C. Leadbeater. 1974. Fine-structural observations on six species of Chrysochromulina from wild Danish marine nanoplankton, including a discription of C. campanulifera sp. nov. and a preliminary summary of the nanoplankton as a whole. Dan. Vidensk. Sel. Biol. Skr. 20:1-26. 83. Markey, D. R. 1974. A possible virus infection in the brown alga Pylaiella littoralis. Protoplasma 80:223-232. 84. Martin, E. L., and R. L. Benson. 1982. Algal viruses, pathogenic bacteria and fungi: introduction and bibliography, p. 793-798. In J. R. Rosowski and B. C. Parker (ed.), Selected papers in phycology II. Phycological Society of America Inc., Lawrence, Kans. 85. Matthews, R. E. F. 1982. Classification and nomenclature of viruses. Intervirology 17:1-199. 86. Mattox, K. R., K. D. Stewart, and G. L. Floyd. 1972. Probable virus infections in four genera of green algae. Can. J. Microbiol. 18:1620-1621. 87. Mayer, J. A., and F. J. R. Taylor. 1979. A virus which lyses the marine nanoflagellate Micromonas pusilla. Nature (London)
281:299-301. 88. McAuley, P. J., and L. Muscatine. 1986. The cell cycle of symbiotic Chlorella. IV. DNA content of algae slowly increases during host starvation of green hydra. J. Cell Sci. 85:73-84. 89. Meier, R., W. Reisser, and W. Wiessner. 1980. Cytological studies on the endosymbiotic unit of Paramecium bursaria Ehbg. and Chlorella spec. II. The regulation of the endosymbiotic algal population as influenced by nutritional condition of the symbiotic partners. Arch. Protistenkd. 123:333-341. 90. Meier, R., and W. Wiessner. 1989. Infection of algae-free Paramecium bursaria with symbiotic Chlorella sp. isolated from green paramecium. II. A timed study. J. Cell Sci. 95:571-579. 91. Meints, R. H. Unpublished results. 92. Meints, R. H., D. E. Burbank, J. L. Van Etten, and D. T. A. Lamport. 1988. Properties of the Chlorella receptor for the virus PBCV-1. Virology 164:15-21. 93. Meints, R. H., K. Lee, D. E. Burbank, and J. L. Van Etten. 1984. Infection of a Chlorella-like alga with the virus, PBCV-1: ultrastructural studies. Virology 138:341-346. 94. Meints, R. H., K. Lee, and J. L. Van Etten. 1986. Assembly site of the virus PBCV-1 in a Chlorella-like green alga: ultrastructural studies. Virology 154:240-245. 95. Meints, R. H., and R. L. Pardy. 1980. Quantitative demonstration of cell surface involvement in a plant-animal symbiosis: lectin inhibition of reassociation. J. Cell Sci. 43:239-251. 96. Meints, R. H., J. L. Van Etten, D. Kuczmarski, K. Lee, and B. Ang. 1981. Viral infection of the symbiotic Chlorella-like alga present in Hydra viridis. Virology 113:698-703. 97. Melkonian, M. 1982. Virus-like particles in the scaly green flagellate Mesostigma viride. Br. Phycol. J. 17:63-68. 98. Mews, L. K. 1980. The green hydra symbiosis. III. The biotrophic transport of carbohydrate from alga to animal. Proc. R. Soc. London Ser. B 209:377-401. 99. Mindich, L., and D. Bamford. 1988. Lipid-containing bacteriophages, p. 475-520. In R. Calendar (ed.), The bacteriophages, vol. 2. Plenum Press, New York. 100. Mitra, A. Personal communication. 101. Moestrup, O., and H. A. Thomsen. 1974. An ultrastructural study of the flagellate Pyramimonas orientalis with particular emphasis on Golgi apparatus activity and the flagellar apparatus. Protoplasma 81:247-269. 102. Moss, B. 1990. Regulation of vaccinia virus transcription. Annu. Rev. Biochem. 59:661-688. 103. Muller, D. G., H. Kawai, B. Stacke, and S. Lanka. 1990. A virus infection in the marine brown alga Ectocarpus siliculosus (Phaeophyceae). Bot. Acta 103:72-82. 104. Muller, D. G., and S. Lanka. Personal communication. 105. Muscatine, L. 1971. Experiments on green algae coexistent with zooxanthellae in sea anemones. Pac. Sci. 25:13-21. 106. Muscatine, L., C. B. Cook, R. L. Pardy, and R. R. Pool. 1975. Uptake, recognition and maintenance of symbiotic Chlorella
by Hydra viridis. Symp. Soc. Exp. Biol. 29:175-203. 107. Nakamura, K., and R. W. Compans. 1979. Host cell- and virus strain-dependent differences in oligosaccharides of hemagglutinin glycoproteins of influenza A viruses. Virology 95:8-23. 108. Narva, K. E., and J. L. Van Etten. Unpublished results. 109. Narva, K. E., J. L. Van Etten, B. E. Slatko, and J. S. Benner. 1988. The amino acid sequence of the eukaryotic DNA [N6adenine]methyltransferase, M.CviBIII, has regions of similarity with the prokaryotic isoschizomer M.TaqI and other DNA [N6-adenine]methyltransferases. Gene 74:253-259. 110. Narva, K. E., D. L. Wendell, M. P. Skrdla, and J. L. Van Etten. 1987. Molecular cloning and characterization of the gene encoding the DNA methyltransferase, M.CviBIII, from Chlorella virus NC-1A. Nucleic Acids Res. 15:9807-9823. 111. Nichols, H. W., and H. C. Bold. 1965. Trichosarcina polymorpha gen. et. sp. nov. J. Phycol. 1:34-38. 112. Nietfeldt, J., K. Lee, and J. L. Van Etten. Chlorella virus PBCV-1 replication is not affected by cytoskeletal disruptors. Intervirology, in press. 113. Nietfeldt, J., and J. L. Van Etten. Unpublished results. 114. Northcote, D. H., K. J. Goulding, and R. W. Horne. 1958. The chemical composition and structure of the cell wall of Chlorella pyrenoidosa. Biochem. J. 70:391-397. 115. O'Brien, T. L., R. MacLeod, and M. C. MacLean. 1984. Absence of lytic virus in two species of symbiotic algae within the sea anemone Anthopleura xanthogrammica (Coelenterata: Anthozoa). Trans. Am. Microsc. Soc. 103:228-232. 116. Oh-Hama, T., and S. Miyachi. 1988. Chlorella, p. 3-26. In M. A. Borowitzka and L. J. Borowitzka (ed.), Micro-algal biotechnology. Cambridge University Press, Cambridge. 117. Oliviera, L., and T. Bisalputra. 1978. A virus infection in the brown alga Sorocarpus uvaeformis (Lyngbye) Pringsheim (Phaeophyta, Ectocarpales). Ann. Bot. 42:439-445. 118. Ortin, J., L. Enjuanes, and E. Vinuela. 1979. Cross-links in African swine fever virus DNA. J. Virol. 31:579-583. 119. Pardy, R. L. Personal communication. 120. Pardy, R. L. 1974. Some factors affecting the growth and distribution of the algal endosymbionts of Hydra viridis. Biol. Bull. 147:105-118. 121. Pardy, R. L. 1983. Preparing aposymbiotic hydra, p. 393-397. In H. M. Lenhoff (ed.), Hydra: research methods. Plenum Press, New York. 122. Pearson, B. R., and R. E. Norris. 1974. Intranuclear virus-like particles in the marine alga Platymonas sp. (Chlorophyta, Prasinophyceae). Phycologia 13:5-9. 123. Pickett-Heaps, J. D. 1972. A possible virus infection in the green alga Oedogonium. J. Phycol. 8:44 47. 124. Pienaar, R. N. 1976. Virus-like particles in three species of phytoplankton from San Juan Island, Washington. Phycologia
15:185-190. 125. Pool, R. R. 1979. The role of algal antigenic determinants in the recognition of potential algal symbionts by cells of chlorohydra. J. Cell Sci. 35:367-379. 126. Posfai, J., A. S. Bhagwat, G. Posfai, and R. J. Roberts. 1989. Predictive motifs derived from cytosine methyltransferases. Nucleic Acids Res. 17:2421-2435. 127. Preisig, H. R., and D. J. Hibberd. 1984. Virus-like particles and endophytic bacteria in Paraphysomonas and Chromophysomonas (Chrysophyceae). Nord. J. Bot. 4:279-285. 128. Que, Q., and J. L. Van Etten. Unpublished results. 129. Reisser, W. 1981. The endosymbiotic unit of Stentor polymorphus and Chlorella sp. Morphological and physiological studies. Protoplasma 105:273-284. 130. Reisser, W. 1986. Endosymbiotic associations of freshwater protozoa and algae. Progr. Protistol. 1:195-214. 131. Reisser, W. 1987. Naturally occurring and artificially established associations of ciliates and algae. Ann. N.Y. Acad. Sci.
503:316-329. 132. Reisser, W., B. Becker, and T. Klein. 1986. Studies on ultrastructure and host range of a Chlorella attacking virus. Protoplasma 135:162-165. 133. Reisser, W., D. E. Burbank, R. H. Meints, B. Becker, and J. L. Van Etten. 1991. Viruses distinguish symbiotic Chlorella spp.
VOL. 55, 1991
of Paramecium bursaria. Endocytobiosis Cell Res. 7:245-257. 134. Reisser, W., D. E. Burbank, S. M. Meints, R. H. Meints, B. Becker, and J. L. Van Etten. 1988. A comparison of viruses infecting two different Chlorella-like green algae. Virology 167:143-149. 135. Reisser, W., D. Fischer-Defoy, J. Staudinger, N. Schilling, and K. Hausmann. 1984. The endosymbiotic unit of Climacostomum virens and Chlorella sp. I. Morphological and physiological studies on the algal partner and its localization in the host cell. Protoplasma 119:93-99. 136. Reisser, W., T. Klein, and B. Becker. 1988. Studies on phycoviruses. I. On the ecology of viruses attacking Chlorellae exsymbiotic from an European strain of Paramecium bursaria. Arch. Hydrobiol. 111:575-583. 137. Richmond, A. (ed.). 1986. Handbook of microalgal mass culture. CRC Press, Inc., Boca Raton, Fla. 138. Rohozinski, J., L. E. Girton, and J. L. Van Etten. 1989. Chlorella viruses contain linear nonpermuted double stranded DNA genomes with covalently closed hairpin ends. Virology 168:363-369. 139. Rohozinski, J., and J. L. Van Etten. 1989. Characterization of DNA polymerases in an uninfected and virus PBCV-1 infected green alga-Chlorella strain NC64A. Intervirology 30:156-162. 140. Rohozinski, J., and J. L. Van Etten. 1989. Purification and characterization of soluble and membrane-associated DNA polymerases from a virus PBCV-1 infected green alga Chlorella NC64A. J. Exp. Bot. 40:1293-1298. 141. Rosowski, J. R., and B. C. Parker (ed.). 1982. Selected papers in phycology II. Phycological Society of America, Inc., Lawrence, Kans. 142. Schlesinger, M. J., and S. Schlesinger. 1987. Domains of virus glycoproteins. Adv. Virus. Res. 33:1-44. 143. Schnepf, E., C. J. Soeder, and E. Hegewald. 1970. Polyhedral virus like particles lysing the aquatic Phycomycete Aphelidium sp., a parasite of the green alga Scenedesmus armatus. Virology 42:482-487. 144. Schuster, A. M., D. E. Burbank, B. Meister, M. P. Skrdla, R. H. Meints, S. Hattman, D. Swinton, and J. L. Van Etten. 1986. Characterization of viruses infecting a eukaryotic Chlorellalike green alga. Virology 150:170-177. 145. Schuster, A. M., L. Girton, D. E. Burbank, and J. L. Van Etten. 1986. Infection of a Chlorella-like alga with the virus PBCV-1: transcriptional studies. Virology 148:181-189. 146. Schuster, A. M., M. Graves, K. Korth, M. Ziegelbein, J. Brumbaugh, D. Grone, and R. H. Meints. 1990. Transcription and sequence studies of a 4.3-kbp fragment from a dsDNA eukaryotic algal virus. Virology 176:515-523. 147. Schuster, A. M., J. A. Waddle, K. Korth, and R. H. Meints. 1990. The chloroplast genome of an exsymbiotic Chlorella-like green alga. Plant Mol. Biol. 14:859-862. 148. Shelef, G., and C. J. Soeder (ed.). 1980. Algal biomass: production and use. Elsevier/North-Holland Biomedical Press, Amsterdam. 149. Sherman, L. A., and R. M. Brown. 1978. Cyanophages and viruses of eukaryotic algae, p. 145-234. In H. Fraenkel-Conrat and R. R. Wagner (ed.), Comprehensive Virology, vol. 12. Plenum Press, New York. 150. Shields, S. L., D. E. Burbank, R. Grabherr, and J. L. Van Etten. 1990. Cloning and sequencing the cytosine methyltransferase gene M.CviJI from Chlorella virus IL-3A. Virology 176:16-24. 151. Shihra, I., and R. W. Krauss. 1965. Chlorella: physiology and taxonomy of forty one isolates. University of Maryland Press, College Park, Md. 152. Sicko-Goad, L., and G. Walker. 1979. Viroplasm and large virus-like particles in the dinoflagellate Gymnodinium uberrimum. Protoplasma 99:203-210. 153. Sieburth, J. M., P. W. Johnson, and P. E. Hargraves. 1988. Ultrastructure and ecology of Aureococcus anophagefferens gen. et sp. nov. (Chrysophyceae): the dominant picoplankter during a bloom in Narragansett bay, Rhode Island, summer 1985. J. Phycol. 24:416-425. 154. Siegel, R. W. 1960. Hereditary endosymbiosis in Paramecium
VIRUSES OF EUKARYOTIC ALGAE
619
bursaria. Exp. Cell Res. 19:239-252. 155. Skotnicki, A., A. Gibbs, and N. G. Wrigley. 1976. Further studies on Chara corrallina virus. Virology 75:457-468. 156. Skrdla, M. P., D. E. Burbank, Y. Xia, R. H. Meints, and J. L. Van Etten. 1984. Structural proteins and lipids in a virus, PBCV-1, which replicates in a Chlorella-like alga. Virology 135:308-315. 157. Skrdla, M. P., M. Graves, J. L. Van Etten, and R. H. Meints. Unpublished results. 158. Slatko, B. E., J. S. Benner, T. Jager-Quinton, L. S. Moran, T. G. Simcox, E. M. Van Cott, and G. G. Wilson. 1987. Cloning, sequencing and expression of the TaqI restrictionmodification system. Nucleic Acids Res. 15:9781-9796. 159. Smith, D. C., and A. E. Douglas. 1987. The biology of symbiosis. Edward Arnold, Ltd., London. 160. Smith, H. O., and S. V. Kelly. 1984. Methylases of the type II restriction-modification systems, p. 39-71. In A. Razin, H. Cedar, and A. D. Riggs (ed.), DNA methylation: biochemistry and biological significance. Springer-Verlag KG, Berlin. 161. Sogo, J. M., J. M. Almendral, A. Talavera, and E. Vinuela. 1984. Terminal and internal inverted repetitions in African swine fever virus DNA. Virology 133:271-275. 162. Soyer, M. 0. 1978. Particules de type viral et filaments trichocystoides chez les Dinoflagelles. Protistologica 14:53-58. 163. Stanker, L. H., and L. R. Hoffman. 1979. A simple histological assay to detect virus-like particles in the green alga Cylindrocapsa (Chlorophyta). Can. J. Bot. 57:838-842. 164. Stanker, L. H., L. R. Hoffman, and R. MacLeod. 1981. Isolation and partial chemical characterization of a virus-like particle from a eukaryotic alga. Virology 114:357-369. 165. Stefan, C., Y. Xia, and J. L. Van Etten. 1991. Molecular cloning and characterization of the gene encoding the adenine methyltransferase M.CviRI from Chlorella virus XZ-6E. Nucleic Acids Res. 19:307-311. 166. Stelker, S., Y. Xia, and J. L. Van Etten. Unpublished results. 167. Strasser, P., Y. Zhang, J. Rohozinski, and J. L. Van Etten. 1991. The termini of the Chlorella virus PBCV-1 genome are identical 2.2-kbp inverted repeats. Virology 180:763-769. 168. Suttle, C. A. Personal communication. 169. Suttle, C. A., A. M. Chan, and M. T. Cottrell. 1990. Infection of phytoplankton by viruses and reduction of primary productivity. Nature (London) 347:467-469. 170. Swale, E. M. F., and J. H. Belcher. 1973. A light and electron microscope study of the colourless flagellate Aulacomonas Skuja. Arch. Mikrobiol. 92:91-103. 171. Takeda, H. 1988. Classification of Chlorella strains by cell wall sugar composition. Phytochemistry 27:3822-3826. 172. Takeda, H. 1988. Classification of Chlorella strains by means of the sugar components of the cell wall. Biochem. Syst. Ecol. 16:367-371. 173. Tessman, I. 1985. Genetic recombination of the DNA plant virus PBCV-1 in a Chlorella-like alga. Virology 145:319-322. 174. Tikhonenko, A. S., and N. B. Zavarzina. 1966. Morphology of the lytic agents of Chlorella pyrenoidosa Mikrobiologiya 35: 850-852. 175. Tomas, R. N., and E. R. Cox. 1973. Observations on the symbiosis of Peridinium balticum and its intracellular alga. I. Ultrastructure. J. Phycol. 9:304-323. 176. Toth, R., and R. T. Wilce. 1972. Viruslike particles in the marine alga Chorda tomentosa Lyngbye (Phaeophyceae). J. Phycol. 8:126-130. 177. Traylor, E., Y. Xia, P. Roy, and J. L. Van Etten. Unpublished results. 178. Trench, R. K. 1979. The cell biology of plant-animal symbiosis. Annu. Rev. Plant Physiol. 30:485-531. 179. Van Etten, J. L. Unpublished results. 180. Van Etten, J. L., D. E. Burbank, J. Joshi, and R. H. Meints. 1984. DNA synthesis in a Chlorella-like alga following infection with the virus PBCV-1. Virology 134:443-449. 181. Van Etten, J. L., D. E. Burbank, D. Kuczmarski, and R. H. Meints. 1983. Virus infection of culturable Chlorella-like algae and development of a plaque assay. Science 219:994-996. 182. Van Etten, J. L., D. E. Burbank, and R. H. Meints. 1986.
620
183. 184. 185. 186.
187.
188. 189.
190.
191.
192.
193. 194. 195. 196.
197. 198. 199.
200. 201. 202.
203. 204.
205.
VAN ETTEN ET AL.
Replication of the algal virus PBCV-1 in UV-irradiated Chlorella. Intervirology 26:115-120. Van Etten, J. L., D. E. Burbank, A. M. Schuster, and R. H. Meints. 1985. Lytic viruses infecting a Chlorella-like alga. Virology 140:135-143. Van Etten, J. L., D. E. Burbank, Y. Xia, and R. H. Meints. 1983. Growth cycle of a virus, PBCV-1, that infects Chlorellalike algae. Virology 126:117-125. Van Etten, J. L., and R. H. Meints. Unpublished results. Van Etten, J. L., R. H. Meints, D. E. Burbank, D. Kuczmarski, D. A. Cuppels, and L. C. Lane. 1981. Isolation and characterization of a virus from the intracellular green alga symbiotic with Hydra viridis. Virology 113:704-711. Van Etten, J. L., R. H. Meints, D. Kuczmarski, D. E. Burbank, and K. Lee. 1982. Viruses of symbiotic Chlorella-like algae isolated from Paramecium bursaria and Hydra viridis. Proc. Natl. Acad. Sci. USA 79:3867-3871. Van Etten, J. L., Q. Que, L. C. Lane, and M. K. Brakke. Unpublished results. Van Etten, J. L., A. M. Schuster, L. Girton, D. E. Burbank, D. Swinton, and S. Hattman. 1985. DNA methylation of viruses infecting a eukaryotic Chlorella-like green alga. Nucleic Acids Res. 13:3471-3478. Van Etten, J. L., A. M. Schuster, and R. H. Meints. 1988. Viruses of eukaryotic Chlorella-like algae, p. 411-428. In Y. Koltin and M. J. Leibowitz (ed.), Viruses of fungi and simple eukaryotes. Marcel Dekker, Inc., New York. Van Etten, J. L., C. H. Van Etten, J. K. Johnson, and D. E. Burbank. 1985. A survey for viruses from freshwater that infect a eukaryotic Chlorella-like green alga. Appl. Environ. Microbiol. 49:1326-1328. Van Etten, J. L., Y. Xia, and R. H. Meints. 1987. Viruses of a Chlorella-like green alga, p. 307-325. In T. Kosuge and E. W. Nester (ed.), Plant-microbe interactions, vol. II. Macmillan Publishing Co., New York. Vivier, E., A. Petitprez, and A. F. Chive. 1967. Observations ultrastructurales sur les Chlorelles symbiotes de Paramecium bursaria. Protistologica 3:325-334. Waddle, J. A., R. H. Meints, D. E. Burbank, and J. L. Van Etten. Unpublished results. Waddle, J. A., A. M. Schuster, K. W. Lee, and R. H. Meints. 1990. The mitochondrial genome of an exsymbiotic Chlorellalike green alga. Plant Mol. Biol. 14:187-195. Wang, I., M. Graves, Y. Li, L. C. Lane, R. H. Meints, and J. L. Van Etten. Unpublished results. Wang, I., L. C. Lane, Y. Li, and J. L. Van Etten. Unpublished results. Wang, I., Q. Que, and J. L. Van Etten. Unpublished results. Waters, R. E., and A. T. Chan. 1982. Micromonas pusilla virus: the virus growth cycle and associated physiological events within the host cells; host range mutation. J. Gen. Virol. 63:199-206. Weis, D. S. 1977. Synchronous development of symbiotic Chlorellae within Paramecium bursaria. Trans. Am. Microsc. Soc. 96:82-86. Weis, D. S. 1978. Correlation of infectivity and concanavalin A agglutinability of algae exsymbiotic from Paramecium bursaria. J. Protozool. 25:366-370. Wiggins, B. A., and M. Alexander. 1985. Minimum bacterial density for bacteriophage replication: implications for significance of bacteriophages in natural ecosystems. Appl. Environ. Microbiol. 49:19-23. Willis, D. B. (ed.). 1985. Iridoviridae. Curr. Top. Microbiol. Immunol. 116:1-173. Willis, D. B., R. Goorha, and A. Granoff. 1984. DNA methyltransferase induced by frog virus 3. J. Virol. 49:86-91. Willis, D. B., and A. Granoff. 1980. Frog virus 3 is heavily
MICROBIOL. REV.
methylated at CpG sequences. Virology 107:250-257. 206. Wilson, G. C. 1988. Type II restriction-modification systems. Trends Genet. 4:314-318. 207. Wittek, R., A. Menna, H. K. Muller, D. Schumperli, P. G. Boseley, and R. Wyler. 1978. Inverted terminal repeats in rabbit poxvirus and vaccinia virus DNA. J. Virol. 28:171-181. 208. Wittek, R., and B. Moss. 1980. Tandem repeats within the inverted terminal repetition of vaccinia virus DNA. Cell 21: 277-284. 209. Xia, Y., D. E. Burbank, L. Uher, D. Rabussay, and J. L. Van Etten. 1986. Restriction endonuclease activity induced by PBCV-1 virus infection of a Chlorella-like green alga. Mol. Cell. Biol. 6:1430-1439. 210. Xia, Y., D. E. Burbank, L. Uher, D. Rabussay, and J. L. Van Etten. 1987. IL-3A virus infection of a Chlorella-like green alga induces a DNA restriction endonuclease with novel sequence specificity. Nucleic Acids Res. 15:6075-6090. 211. Xia, Y., D. E. Burbank, and J. L. Van Etten. 1986. Restriction endonuclease activity induced by NC-1A virus infection of a Chlorella-like green alga. Nucleic Acids Res. 14:6017-6030. 212. Xia, Y., R. Morgan, I. Schildkraut, and J. L. Van Etten. 1988. A site-specific single strand endonuclease activity induced by NYs-1 virus infection of a Chlorella-like green alga. Nucleic Acids Res. 16:9477-9487. 213. Xia, Y., K. E. Narva, and J. L. Van Etten. 1987. The cleavage site of the RsaI isoschizomer, CviII, is G/ITAC. Nucleic Acids Res. 15:10063. 214. Xia, Y., M. P. Skrdla, and J. L. Van Etten. Unpublished
results. 215. Xia, Y., and J. L. Van Etten. Unpublished results. 216. Xia, Y., and J. L. Van Etten. 1986. DNA methyltransferase induced by PBCV-1 virus infection of a Chlorella-like green alga. Mol. Cell. Biol. 6:1440-1445. 217. Yamada, T. 1982. Isolation and characterization of chloroplast DNA from Chlorella ellipsoidea. Plant Physiol. 70:92-96. 218. Yamada, T. 1983. Characterization of inverted repeat sequences and ribosomal RNA genes of chloroplast DNA from Chlorella ellipsoidea. Curr. Genet. 7:481-487. 219. Yamada, T., and K. Sakaguchi. 1982. Comparative studies on Chlorella cell walls: induction of protoplast formation. Arch. Microbiol. 132:10-13. 220. Yamada, T., and K. Sakaguchi. 1982. Electron microscopic studies of Chlorella ellipsoidea protoplast formation. J. Gen. Microbiol. 128:1319-1327. 221. Yao, M. C., and C. H. Yao. 1981. Repeated hexanucleotide C-C-C-C-A-A is present near free ends of macronuclear DNA of Tetrahymena. Proc. Natl. Acad. Sci. USA 78:7436-7439. 222. Yonker, C. R., K. D. Caldwell, J. C. Giddings, and J. L. Van Etten. 1985. Physical characterization of PBCV virus by sedimentation field flow fractionation. J. Virol. Methods 11:145160. 223. Yuen, L., and B. Moss. 1987. Oligonucleotide sequence signaling transcriptional termination of vaccinia virus early genes. Proc. Natl. Acad. Sci. USA 84:6417-6421. 224. Zavarzina, N. B. 1961. A lytic agent in cultures of Chlorella pyrenoidosa Pringh. Dokl. Akad. Nauk SSSR Ser. Biol. 137: 435-437. 225. Zavarzina, N. B. 1964. Lysis of Chlorella cultures in the absence of bacteria. Mikrobiologiya 33:561-564. 226. Zavarzina, N. B., and A. E. Protsenko. 1958. The lysis of Chlorella pyrenoidosa Pringh cultures. Dokl. Akad. Nauk SSSR Ser. Biol. 122:936-939. 227. Zhang, Y., D. E. Burbank, and J. L. Van Etten. 1988. Chlorella viruses isolated in China. Appl. Environ. Microbiol. 54:21702173. 228. Zhang, Y., and J. L. Van Etten. Unpublished results.
